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C H A P T E R 6
Examples of Pathway
Manipulations: Metabolic
Engineering in Practice
Nature has provided a remarkable array of metabolic pathways as wit-
nessed by the diversity of extant microorganisms. In certain cases, the
assembly and kinetic coordination of such pathways in a particular organism
are suitable for a useful commercial application. In most cases, however,
genetic improvements are required for the optimization of the conversion
reactions and kinetic properties of a cell to render it suitable for practical
use. These improvements are guided by the current understanding of micro-
bial metabolism and molecular genetics and implemented by molecular
biological techniques and recombinant DNA technology. The rational trans-
fer of conversion pathways has produced new and desirable functionalities in
cells, thus benefiting the pharmaceutical, agricultural, food, chemical, and
environmental sectors.
In this chapter we review applications of metabolic pathway manipulation.
We follow the classification of Cameron and Tong (1993), slightly modified,
in organizing the large number of examples in basically five groups, i.e.,
203
204 Metabolic Engineering
applications aiming at (a) yield and productivity improvement of products
made by microorganisms, (b) expansion of the range of substances that can
be metabolized by an organism, (c) formation of new and novel products,
(d) general improvement of cellular properties, and (e) xenobiotic degrada-
tion. We have three goals in reviewing these applications in the context of
this book. First, we provide a sample of the truly enormous range of
possibilities for biocatalyst improvement afforded by pathway manipulation
and metabolic engineering. We note that this review is focused, almost
exclusively, on industrial applications. Except for the introduction (Chapter
1) and a few passing references, very little is included on the broad
applications of metabolic engineering in the medical field. The reason is,
simply, that most of the work in this field is current and not sufficiently
crystallized for our review purposes. Reported progress, however, leaves little
doubt about the impact of the tools and methodologies of metabolic engi-
neering in analyzing tissue and organs in vivo and in vitro, as well as
providing fundamentals for the rational analysis of the organization and
function of signal transduction pathways.
The second goal of this review is to provide the reader with a sense of the
complexity of metabolic pathways, along with their regulation and coordina-
tion with the overall metabolism. An important corollary of the admittedly
complex structure of these pathways is that the methods for their systematic
analysis may not always be as simple as one might desire. This point will
become clearer as we delve into the methods of metabolic flux determination
(Chapters 8 and 9) and issues of control and flux amplification in complex
metabolic networks (Chapters 11 and 12). In the same vein, representative
assays of the state of cell metabolism, metabolic production pathways, or
signal transduction pathways may require a multidimensional array of mea-
surements, in contrast to current practice. The recognition of truly distin-
guishing pattems in large volumes of measurements will be a challenge, and
methods of flux analysis could provide a useful framework to this end.
The third goal of this review is to underscore the methods used for
effecting desired changes in cellular systems for industrial use or medical
reasons. It will become apparent that most successful applications require
coordinated modification of more than one enzymatic step in a metabolic
network. This is almost necessary for those applications that extend beyond a
simple product-forming pathway and involve the complex structures of
central carbon metabolism. We believe that this will become a generic
requirement as research focus gradually shifts from the simplicity of highly
reduced model systems to the realm of realistic industrial or medical situa-
tions. As with complex measurement interpretation mentioned earlier,
metabolic engineering can have an impact in the rational design of metabolic
pathway modifications.
6 Examples of Pathway Manipulations 205
6.1. ENHANCEMENT OF PRODUCT YIELD
AND PRODUCTIVITY
A large number of mainly industrial applications can be classified in this
group. We note that, although often interchangeable in a loose sense, yield
and productivity represent different figures of merit that also require differ-
ent strategies for their enhancement. Yield impacts primarily the cost of raw
materials and is affected by redirection of metabolic fluxes toward the forma-
tion of the desired product. Productivity, on the other hand, is the key
determinant of the capital cost of bioprocessing equipment and can be
improved by amplification of metabolic fluxes. Admittedly, overall process
optimization must include both yield and productivity concerns, although in
certain cases decoupling of the two may be possible. Productivity depends,
first and foremost, on the specific rate of substrate uptake, which for most
industrial organisms ranges between 0.2 and 0.5 of substrate per gram of
biomass per hour. If such uptake rates are realized under process conditions,
then productivity can be economically acceptable provided that byproduct
formation is minimized. Under these conditions, yield and productivity
optimization methods may indeed converge. If, on the other hand, uptake
rates are too low, then productivity optimization should begin with the
amplification of the substrate transport system, followed by flux redirection
as dictated by strategies of yield optimization.
Yield and productivities obviously are more important in large volume,
low cost industrial operations. We next review efforts aimed at the improve-
ment of yield and productivity of ethanol, amino acids, and solvents by
metabolic engineering.
6.1.1. ETHANOL
Ethanol is an important industrial chemical with emerging potential as a
biofuel to replace vanishing fossil fuels. Additionally, it may have significant
environmental impact as ethanol combustion is less polluting, and it may
serve as feedstock for the production of oxygenated fuels. Also, because its
production is mainly based on agricultural products, it will enhance the
"carbon cycle" for atmospheric CO 2 removal. According to current estimates,
the United States will be importing more than 50% of its crude oil and
refined products to meet energy needs in the year 2010. From an economic
point of view, ethanol (and other biofuels), by providing domestic resources
to meet part of this demand, can play a major role in stabilizing energy
206 Metabolic Engineering
prices, improving national energy security, and ensuring rural and regional
economic development.
Ethanol can be made from a number of renewable feedstocks, including
sugar crops such as sugar-cane, starch-containing grains such as corn, or
lignocellulosic materials including agricultural residues, herbaceous crops,
and wood. The economics of the ethanol process is determined by the cost of
sugar. Almost all of the U.S. fuel ethanol production of 2.3 billion liters was
made from corn, and it is estimated that an additional 20 billion liters of
ethanol per year could be made with surplus corn. Over the past decade, the
cost of ethanol has dropped from more than $1.0 L -I to approximately
$0.3-0.5 L -1, with a projected cost of less than $0.25 L -1 in the near future.
Lignocellulosic materials are such an abundant and inexpensive resource
that existing supplies could support the sustainable production of liquid
transportation fuels on the same scale as the total US consumption. The
National Renewable Energy Laboratory (NREL) hasestimated the current
cost of ethanol production from lignocellulose to be about $0.32 L -1, assum-
ing a feedstock cost of $42 per dry ton (National Renewable Energy Labora-
tory, 1996). This average biomass cost amounts to approximately $0.06 kg -1
of sugar or a contribution to the feedstock costs for ethanol production of as
low as $0.10 L -1. Lignocellulosic crops considered to be suitable raw materi-
als for fuel ethanol production are fast-growing wood, agricultural and
forestry residues and various kinds of wastes, e.g., pulping waste, newsprint,
and municipal solid waste. Efficient utilization of the hemicellulose compo-
nent of lignocellulosic feedstocks (25 % dry weight of hardwood and predom-
inantly D-xylose) offers an opportunity to reduce the cost of producing fuel
ethanol by 25% (Bull, 1990). Whereas lignocellulose is inexpensive because
it cannot be digested and therefore does not compete as a food, its inability
to be digested also makes it difficult to convert to fermentable sugars.
Furthermore, lignocellulose is a complex structure with three major compo-
nents (cellulose, hemicellulose, and lignin), each of which must be processed
separately to make the best use of the high efficiencies inherent in biological
processes. A general process schematic for the conversion of lignocellulose to
ethanol is shown in Fig. 6.1. The hydrolysate, resulting after prehydrolysis
and hydrolysis, contains varying amounts of monosaccharides, in the form of
both pentoses (D-xylose and L-arabinose) and hexoses (Table 6.1) and a
broad range of substances either derived from the raw material or formed as
reaction byproducts from the pretreatment stage of the process (sugar and
lignin degradation products). Xylose is the most abundant sugar in the
hemicellulose of hardwoods and crop residues, whereas mannose is more
abundant in the hemicellulose of softwoods. Furthermore, xylose is second
only to glucose in natural abundance. Microbial conversion of the sugar
residues present in wastepaper and yard trash from U.S. landfills alone could
6 Examples of Pathway Manipulations 207
T ~,d,ydro,ys,~)
1
I~,o,o~o I I• T I ,se I
! L,qo,o ~o.~ I.T.A.oL I
FIGURE 6.1 Conversion of lignocellulose to ethanol. Crystalline cellulose, the largest (50%)
and most difficult fraction, is hydrolyzed by a combination of acid and enzymatic processes.
During these steps 95-98% of the xylose and glucose is recovered. These monosaccharides
subsequently are converted to ethanol by appropriate microorganisms.
TABLE 6.1 Carbohydrate Structural Polymers in Lignocellulose
for a Typical Softwood, Such as Common Beech
Polymer Monomer(s) Typical % Total
Cellulose Glucose 40
Hemicellulose Xylose 30
Arabinose
Mannose
Glucose
Galactose
Lignin Phenylpropane 25
Pectin Uronic acids 5
208 Metabolic Engineering
provide more than 400 billion liters of ethanol (Lynd et al., 1991), 10 times
the corn-derived ethanol burned annually as a 10% blend with gasoline
(Keim and Venkatasubramanian, 1989).
The fermentation organism must be able to ferment all monosaccharides
present and, in addition, withstand potential inhibitors in the hydrolysate.
The most commonly used ethanol producer, Saccharomyces cerevisiae, cannot
ferment pentoses, which may constitute 8-28% of the raw material (Ladisch
et al., 1983). Yeasts produce ethanol efficiently from hexoses by the pyruvate
decarboxylase-alcohol dehydrogenase (PDC-ADH) system. However, during
xylose fermentation the byproduct xylitol accumulates, thereby reducing the
yield of ethanol. Furthermore, yeasts are reported to ferment L-arabinose
only very weakly. The efficient fermentation of xylose and other hemicellu-
lose constituents may prove essential for the development of an economically
viable process to produce ethanol from biomass.
Pentose-fermenting microorganisms are found among bacteria, yeasts, and
fungi, with the yeasts Pichia stipitis, Candida shehatae, and Pachysolen
tannophilus being the most promising naturally occurring microorganisms.
Only a handful of bacterial species are known that do possess the important
PDC-ADH pathway to ethanol. Among these, Zymomonas mobilis has the
most active PDC-ADH system; however, it is incapable of dissimilating
pentose sugars. During recent years, the application of metabolic engineering
resulted in recombinant bacteria (Alterhum and Ingram, 1989; Feldmann
et al., 1989; Ingram et al., 1987; Ohta et al., 1991a,b; Tolan and Finn,
1987a,b) and yeasts (Hallborn et al., 1991; K/fitter et al., 1990; Tantirungkij
et al., 1993) as competent ethanol producers. A recent study that compared
the performance of various ethanol producers, natural and recombinant, in
pentose-rich com cob hydrolysate concluded that the recombinant ethanolo-
genic Escherichia coli KOll (E. coli carrying Z. mobilis pdc and adhB
integrated on the chromosome) is currently the best fermentation organism
(Hahn-H~igerdal et al., 1993).
Initial studies were only partially successful in redirecting fermentative
metabolism in Erwinia chrysanthemi (Tolan and Finn, 1987a,b), Klebsiella
planticola (Tolan and Finn, 1987a,b) and E. coli (Brau and Sahm, 1986). The
first generation of recombinant organisms amplified the PDC activity only
and depended on endogenous levels of ADH activity to couple the further
reduction of acetaldehyde to the oxidation of NADH (see Fig. 6.2). Because
ethanol is just one of a number of fermentation products normally produced
by these enteric bacteria, a deficiency in ADH activity together with NADH
accumulation contributed to the formation of various unwanted byproducts.
This problem was solved by amplifying the ADH activity through overexpres-
sion of the Z. mobilis adhB gene yielding recombinants of E. coli (Ingram
et al., 1987) and K. oxytoca (Ohta et al., 1991a,b; Wood and Ingram, 1992)
6 Examples of Pathway Manipulations 209
Glucose
I
EMP
/ ~mvate ~ P D N - ~ T C A
POC JPFL LDH
ate 1
FIGURE 6.2 Competing pathways at the pyruvate branch point. Abbreviations: EMP, Emb-
den-Meyerhof-Parnas enzymes and intermediates; PDC, pyruvate decarboxylase; ADH, alcohol
dehydrogenase; PFL, pyruvate formate-lyase; ACK/PTA, phosphotransacetylase and acetate
kinase; ALDH, acetaldehyde dehydrogenase; FHL, formate hydrogen lyase; LDH, lactate dehy-
drogenase; PDH, pyruvate dehydrogenase.
that efficiently ferment a variety of sugars to ethanol. This was accomplished
by assembling both Z. mobilis genes (i.e., pdc and adhB) into an artificial
operon to produce a portable genetic element for ethanol production (PET
operon). E. coli is an advantageous host organism, especially because it can
grow efficiently on a wide range of carbon substrates that includes five-carbon
sugars.
Pyruvate decarboxylase catalyzes the nonoxidative decarboxylation of
pyruvate to produce acetaldehyde and carbon dioxide (Fig. 6.2). Two alcohol
dehydrogenase isozymes are present in E. coli that catalyze the reduction of
acetaldehyde to ethanol during fermentation accompanied by the oxidation
of NADH to NAD +. In the recombinant E. coli, both enzymes [pyruvate
decarboxylase, (PDC), and alcohol dehydrogenase (ADH)], required to divert
pyruvate metabolism to ethanol are present at high levels. The combined
effect of high PDC levels and low apparent K m (Table 6.2) of this enzyme for
pyruvate effectively is to divert carbon flow to ethanol even in the presence
of native fermentation enzymes like lactate dehydrogenase.
210 Metabolic Engineering
TABLE 6.2 Comparison of Apparent K m Values for Pyruvate for Selected
E. coli and Z. mobilis Pyruvate-Acting Enzymes a
Km
Organism Enzyme Pyruvate NADH
E. coli PDH 0.4 mM 0.18 mM
LDH 7.2 mM > 0.5 mM
PFL 2.0 mM
ALDH 50 ~M
NADH-OX 50 ~M
Z. mobilis PDC 0.4 mM
ADHII 12 ~M
a Abbreviations: PDH, pyruvate dehydrogenase; LDH, lactate dehydrogenase;
PFL, pyruvate formate lyase; ALDH, aldehyde dehydrogenase; NADH-OX,
NADH oxidase; PDC, pyruvate decarboxylase; ADH II, alcohol dehydrogen-
ase II.
Significant amounts of ethanol were produced in recombinant E. coli
containing the pet operon under both aerobic and anaerobic conditions
(Table 6.3). Under aerobic conditions, wild-type E. coli metabolizes pyruvate
through PDH and PFL (Km 0.4 and 2.0 mM, respectively, Table 6.3), with
main products CO 2 and acetate (formed by the conversion of excess acetyl-
CoA). The apparent K m for the Z. mobilis PDC is similar to that of PDH and
lower than those of PFL and LDH, thereby facilitating acetaldehyde produc-
tion. NAD + regeneration under aerobic conditions primarily results from
biosynthesis and from the NADH oxidase (coupled to the electron transport
system). Again, because the apparent K m for Z. mobilis ADH II is over 4-fold
lower than that for E. coli NADH oxidase, the heterologous ADH II effec-
tively competes for endogenous pools of NADH, allowing the reduction of
TABLE 6.3 Comparison of Fermentation Products during Aerobic and Anaerobic
Growth of Wild-Type and Recombinant E. coli a
Fermentation Product (mM)
Growth Plasmid Ethanol Lactate Acetate Succinate
Aerobic
Anaerobic
None 0 0.6 55 0.2
PLO 1308-10 (PET) 337 1.1 17 4.9
None 0.4 22 7 0.9
PLO1308-10 (PET) 482 10 1.2 5.0
a From Ingram and Conway, 1988.
6 Examples of Pathway Manipulations 211
acetaldehyde to ethanol. Under anaerobic conditions, wild-type E. coli me-
tabolizes pyruvate primarily via LDH and PFL. As indicated again in Table
6.2, the apparent K m values for these two enzymes are 18-fold and 5-fold
higher, respectively, than that for Z. mobilis PDC. Furthermore, the apparent
K m values for primary native enzymes involved in NAD + regeneration are
also considerably higher in E. coli than those of Z. mobilis ADH. Overall,
overexpressed ethanologenic Z. mobilis enzymes in E. coli are quite competi-
tive with respect to the native enzymes in channeling carbon (pyruvate) and
reducing power (NADH) into ethanol.
In March 1991, the University of Florida was awarded U.S. Patent No.
5,000,000 for the ingenious microbe created at its Institute of Food and
Agricultural Sciences. The fermentation characteristics of the recombinant
E. coli strain have been reported in numerous studies. Typical final ethanol
concentrations are in excess of 50 g L -1 [e.g., 54.4. and 41.6 g L -1 w e r e
obtained from 10% glucose and 8% xylose, respectively (Ohta et al.,
1991a,b)] at nearmaximum theoretical yields of 0.5 g of ethanol/g of sugar
(sugar ~ 2ethanol + 2CO2). Published volumetric and specific ethanol pro-
ductivities with xylose in simple batch fermentations are 0.6 g of ethanol
(L h) -1 and 1.3 g of ethanol (g DW h) -1, respectively (Alterhum and Ingram,
1989). Further improvements have resulted in volumetric productivities of as
high as 1.8 g of ethanol (L h) -1 (Ohta et al., 1991). The production cost of
ethanol from pentoses (e.g., willow or pine) using E. coli KOll is estimated
at around $0.13 L -1 ( y o n Sivers and Zacchi, 1995; von Sivers et al., 1994),
which can easily bring the final cost of ethanol well below the $0.18 L -1
target for the year 2000. In addition, this ethanologenic E. coli also has the
ability to ferment- besides xylose - all other sugar constituents of lignocellu-
losic material: glucose, mannose, arabinose, and galactose. When the recom-
binant strain was grown on mixtures of sugars typically present in hemicellu-
lose hydrolysates, sequential utilization was observed with glucose consumed
first, followed by arabinose and xylose, to produce near-maximum theoreti-
cal yields of ethanol (Takahashi et al., 1994).
Recently, Ohta et al. investigated the expression of the pdc and adh genes
of Z. mobilis in a related enteric bacterium, Klebsiella oxytoca (Ohta et al.,
1991a,b). In Klebsiella strains, two additional fermentation pathways are
present compared with E. coli (Fig. 6.2), converting pyruvate to succinate
and butanediol. As in the case of E. coli, it was possible to divert more than
90% of the carbon flow from sugar catabolism away from the native
fermentative pathways and toward ethanol. Overexpression of recombinant
PDC alone produced only about twice the ethanol level of the parental strain.
However, when both PDC and ADH were elevated in K. oxytoca M5A1,
ethanol production was both very rapid and efficient: volumetric productivi-
212 Metabolic Engineering
ties ) 2.0 g (L h) -1, yields 0.5 g of ethanol/g of sugar, and final ethanol of
45 g L "1 for both glucose and xylose carbon sources were obtained.
6.1.2. AMINO ACIDS
Amino acids have a wide spectrum of commercial use as food additives, feed
supplements, infusion compounds, therapeutic agents, and precursors for the
synthesis of peptides or agrochemicals. Most microorganisms have the
metabolic machinery to synthesize all essential amino acids from carbon and
nitrogen sources (Fig. 6.3). It is also possible that certain microorganisms can
overproduce one or a group of amino acids. In the mid-1950s, for example,
Japanese scientists isolated a novel bacterium that excreted large quantities of
L-glutamate, giving rise to a new era of amino acid production by fermenta-
tion (Kinoshita et al., 1957). Before that the main sources of amino acids had
been natural proteins and, to a lesser extent, chemical synthesis. This
bacterium, later known as Corynebacterium glutamicum, is a Gram-positive,
short aerobic rod capable of excreting very large amounts of glutamate into
the medium, close to 100 g L -1 under certain conditions.
The success in the industrial production of glutamic acid stimulated
further interest in isolating producing strains for other amino acids. Wild-type
strains of glutamic acid bacteria are capable of producing only a few amino
acids extracellularly, such as glutamic acid, valine, proline, glutamine, and
alanine. For the extracellular accumulation of a desired amino acid, changes
in the cellular metabolism and/or regulatory controls are required. For many
years following the discovery of these bacteria, attempts were made to induce
auxotrophic and regulatory mutants (see Example 5.1). The rationale of
utilizing auxotrophic mutants is to bypass feedback control (see Chapter 5)
by minimizing the intracellular accumulation of feedback inhibitors or
repressors or by modifying the inhibitor binding site, thus rendering the
enzyme insensitive to the presence of the inhibitor. For example, an or-
nithine producer was isolated by using an arginine auxotrophic mutant
followed, a year later, by homoserine auxotrophic mutants for successful
lysine fermentation (Kinoshita et al., 1957). Most amino acids are produced
today by use of strains that contain combinations of auxotrophic and
regulatory mutations. More than 500,000 tons of L-glutamate are produced
annually with C. glutamicum, whereas while its auxotrophic mutant is
responsible for about 400,000 tons of L-lysine per year. The demand for
amino acids is constantly increasing.
In light of the commercial importance of the aspartate family of amino
acids, particularly lysine, intense strain improvement programs were carried
out to isolate strains with superior properties. These programs initially
6 Examples of Pathway Manipulations 213
/ . ,~Glucose (C6)j~
Phenylalanine ( g N ) ~ _ ~ ~ I Triose 1 (C3) I . . . . I
Tyrosine (CgN3) ~ Shikimate (C7)
Tryptophan (C 11N2)
Alanine (C3N) ~ -
Lysine (C6N2)
I
Aspartate (C4N)
Threonine (C4N)
Isoleucine (C6N)
I Pyruvate (C3) H (C5) 1 I
l//
Oxaloacetate (C4) !
Methionine (CsNS)
~x-Ketoglutarate (Cs)
Glycine (C2N)
+= Serine (C3N)
Cysteine (C3NS)
Valine (CsN)
Leucine (CeN)
Citrate (06) [
Proline (CsN)
Glutamate (CsN)
Arginine (C6N4)
FIGURE 6.3 Amino acid biosynthesis from glucose.
employed random mutation and auxotrophic selection procedures utilizing
the wellunderstood metabolic pathways of amino acid biosynthesis and
regulation. Vqild-type Corynebacteria do not accumulate lysine due to con-
certed feedback inhibition of aspartokinase by threonine and lysine (see also
Section 10.1.1 and Fig. 10.1). Thus, the first lysine-producing mutant was a
homoserine dehydrogenase (HDH) deficient and, hence, homoserine-aux-
otrophic strain; incapable of homoserine synthesis and, therefore, requiring
homoserine or threonine and methionine supplementation in the medium.
The latter is provided at a controlled rate so as to satisfy protein synthesis
needs and allow lysine to accumulate to high concentrations at the same
214 Metabolic Engineering
time. Further improvement involved strains resistant to S-(2-aminoethyl)-L-
cysteine (AEC), a lysine analogue. Because AEC resembles lysine, it elicits
similar inhibitory effects, such as inhibition of aspartokinase and arrest of
lysine synthesis. AEC-resistant strains apparently involve deregulated aspar-
tokinases that are not inhibited by lysine and, as such, can accumulate large
amounts of lysine in the medium. Subsequent efforts focused on central
carbon metabolism in an attempt to divert increased amounts of carbon away
from the respiratory and into the anaplerotic pathways. For this purpose,
mutants with citrate-synthase attenuated activity were isolated and found to
offer further improvement in lysine yield, especially in combination with the
previous two phenotypes of HDH deficiency and AEC resistance. This theme
has since been applied with many variations, leading to fluoropyruvate-sen-
sitive (i.e., pyruvate dehydrogenase-attenuated) mutants, alanine auxotrophs,
and many others. The exact nature of the mechanism(s) responsible for any
claimed improvements in these strains is not known due to the poor
characterization of random mutations and also the incomplete understanding
of the function and regulation of the anaplerotic pathways in C. glutamicum.
In recent years, even more potent producing strains have been obtained by
further pathway manipulation, e.g. by eliminating the ability of the produc-
tion strain to degrade the product or by improving cell permeability to favor
excretion of the final product. Several research groups have independently
initiated research programs focusing on the development of metabolic engi-
neering tools for Corynebacterium species. Essential prerequisites are the
availability of vectors derived from endogeneous plasmids and efficient DNA
transfer systems. Small, cryptic plasmids were isolated in various Corynebac-
terium strains, and new and efficient transformation techniques have been
developed in the past few years. This facilitated the isolation of amino acid
biosynthetic genes from Corynebacteria, which currently number around 50
or so [see the review by Jetten and Sinskey (1995)]. These genes, either
individually or in combination, can be utilized to improve production strains
by raising the activities of enzymes or by removing the feedback regulation of
critical enzymes. Use of these genes also allows for specific probes to be
utilized in order to elucidate the biochemistry of amino acid synthesis and
central carbon metabolism (CCM) in general.
Tryptophan
Tryptophan synthesis in E. coli is highly regulated by a complex set of
feedback mechanisms. By transducing each mutation one at a time, re-
searchers combined a long list of alterations to these mechanisms within a
single strain, thus creating a tryptophan overproducer (Aiba et al., 1980;
Shio, 1986). The first step of the aromatic pathway, the conversion of
6 Examples of Pathway Manipulations 215
erythrose-4-P and PEP to 3-deoxy-D-arabino-heptulosonate-7-P (DAHP), is
catalyzed by three isofunctional enzymes (AroF, AroG, and AroH) regulated
by tyrosine, tryptophan, and phenylalanine, respectively. One of the initial
approaches was to simplify the regulation of the system by deleting aroG and
aroH. Furthermore, the tyrosine-regulated enzyme (AroF) was rendered
insensitive to feedback inhibition by mutation (aroF394), and the repression
of this gene was removed by inactivating the repressor gene (tyrR). Other
modifications included the removal of branches leading to tyrosine and
phenylalanine (tyrA and pheA), inactivation of the gene for tryptophanase
(tna) to prevent the possible degradation of the synthesized tryptophan,
alleviation of the feedback inhibition of the tryptophan branch by making
anthranilate synthetase insensitive to tryptophan (trpE382), inactivation of
the tryptophan repressor (trpR); and destruction of the cell's attenuation
control by mutating the gene for tryptophanyl-tRNA synthetase (trpS). The
industrial E. coli strain (NST100) produces about 6.2 g L -1 tryptophan when
cultured in a medium containing 5% glucose for 24 h. Higher tryptophan
yields are possible with the addition of anthranilate to the cultivation
medium.
Recently, a C. glutamicum strain able to produce 18 g L -1 tryptophan has
been altered to produce large amounts of tyrosine (26 g L -1) by overexpress-
ing deregulated 3-deoxy-D-arabino-heptulosonate-7-phosphate (DAHP) syn-
thase and chorismate mutase (Ikeda and Katsumata, 1992). Overexpression
of an additional gene in the previous construct, prephenate dehydratase, led
to the predominant production of phenylalanine (28 g L -1).
Alanine
Uhlenbusch et al. were able to construct a Z. mobilis alanine overproducer
by introducing the gene for alanine dehydrogenase (alaD) from Bacillus
sphaericus (Uhlenbusch et al., 1991). Alanine yield reached 10 mmol per
280 mmol of glucose, which was later increased to 41 mmol by the addition
of 85 mM ammonia that was apparently limiting before. At this production
rate growth ceased, presumably due to the strong competition for pyruvate
between pyruvate decarboxylase (PDC) and alanine dehydrogenase. Starva-
tion for the PDC cofactor thiamine-PP resulted in further growth inhibition
and higher alanine yields (84 mmol in 25 h).
Threonine
Whereas lysine and methionine can be manufactured economically for use
as feed additives, the demand for threonine cannot yet be filled due to the
low yields of existing processes. Recently, however, significant progress was
216 Metabolic Engineering
i Eq/throse_4.p [ DAHP Syntha~
__ ~AroF-*
+ --AroG--~ D~
Indole +
Pyruvate +
NH 3
Chorismate J ) ,
Anthanylate Chodsmate
Synthase (trpE) Mutase
/ ,
I Prephanate
J "
Prephanate
Dehydratase
.~,.,
Prephanate
Dehydrogenase-"~
p-Hydroxyphenylpyruvate
FIGURE 6.4 Aromatic acid biosynthesis. In E. coli, the structural genes form an operon
(trpEDCBA) under a common operator. The regulator gene, which is situated away from this
operon, allows for feedback inhibition of enzyme formation (repression) by the end product
tryptophan.
made in the efforts for the construction of efficient threonine-producing
strains by metabolic engineering. The C. glutamicum genes encoding the first
two enzymes in the threonine pathway, homoserine dehydrogenase (HD) and
homoserine kinase (HK), were isolated by complementation of the E. coli
thrB mutant (Follettie et al., 1988). These two genes form an operon that is
expressed from a single promoter upstream of the hom gene (Peoples et al.,
1988), which is regulated at the transcriptional level by methionine via a
unique attenuation system (Jetten et al., 1993). The final step in the
threonine pathway involves the conversion of homoserine phosphate to
threonine by the constitutive enzymethreonine synthase (TS). The gene
6 Examples of Pathway Manipulations 217
encoding TS ( thrC) was obtained recently by complementation of a C.
glutamicum auxotroph (Han et al., 1990).
Threonine production is determined by the distribution of metabolic
fluxes at the common substrate aspartate-/3-semialdehyde (ASA) to the diver-
gent threonine and lysine biosynthetic pathways (Fig. 6.5). This flux distribu-
tion is controlled by the relative affinities of the competing enzymes, ho-
moserine dehydrogenase and dihydropicolinate synthase, for the common
ppc
asd ~ horn
ddh
. . . . . . . . i
L,0,uco ;1
l
-...
22/
I .......... # , , o , n . }
ilvABNC [ -- ~J~ Isoleucine~
lysA (',,'n.}
FIGURE 6.5 Biosynthetic pathways for aspartate amino acids. Abbreviations: ppc, PEP carbox-
ylase; pc, pyruvate carboxylase; pk, pyruvate kinase; ask, aspartokinase; asd, ASA dehydroge-
nase; dapA, DHP synthase; dapB, DHP reductase; ddh, DAP dehydrogenase; lysA, DAP decar-
boxylase; hom, homoserine dehydrogenase; thrB, homoserine kinase; thrC, threonine synthase;
ilvA, threonine dehydratase (deaminase); ilvBN, acetohydroxy acid synthase, acetohydroxy acid
isomeroreductase, dihydroxy acid dehydratase; ilvC, transaminase C.
218 Metabolic Engineering
substrate ASA. The activity of homoserine dehydrogenase is highly sensitive
to allosteric inhibition by L-threonine (K i = 0.16 mM), and, therefore, under
nominal growth conditions, ASA predominantly enters the lysine pathway.
Critical to the understanding of the molecular basis of threonine inhibition
of homoserine dehydrogenase, as well as to the construction of threonine
overproducers, has been the isolation of feedback-resistant HD (HDar). Two
groups have independently isolated and characterized genes encoding dereg-
ulated, feedback-resistant homoserine dehydrogenase (Archer et al., 1991;
Reinscheid et al., 1991). Archer et al. (1991) determined that the hom ar
mutation is due to a single base deletion that radically alters the structure of
the carboxy terminus, leading to 10 amino acid changes and deletion of the
last 7 residues relative to the wild type. These residues apparently are part of
the threonine binding site, and their removal from the HD ar mutant renders
the HD activity insensitive to feedback inhibition by threonine.
In order to study the regulation of carbon fluxes around the ASA node,
well-defined recombinant strains were constructed (Col6n et al., 1993;
Eikmanns et al., 1991; Reinscheid et al., 1994). Amplification of the threo-
nine genes in wild-type C. g lu tamicum 13032 did not yield any threonine
secretion, presumably due to the feedback inhibition of aspartokinase and
homoserine dehydrogenase by threonine (Eikmanns et al., 1991). Amplifica-
tion of the deregulated horn ar alone (plasmid pJD4, Table 6.4) yielded an
approximately equal flux split between the lysine and threonine pathways,
along with intracellular accumulation of threonine (100 mM) and homoser-
ine (74 mM), which led to the conclusion that threonine production is
probably limited by either its efflux a n d / o r a possible lack of balance
between the activities of HD and HK. In order to prevent an intracellular
buildup of homoserine, Col6n et al. fused the thrB gene to the tac promoter
(plasmid pGC42, Table 6.4) and regulated homar/Ptac- thrB expression by
TABLE 6.4 Threonine Production by C. glutamicum Recombinant Strains a
Excreted amino acids (g L 1)
C. glutamicum strain Lysine Threonine Homoserine Glycine Isoleucine
ATCC 21799(pM2) b 22.0 • 1.0 ( 0.1 ( 0.1 ( 0.1 ~ 0.1
ATCC 21799(pJD4) 4.5 • 0.2 5.4 • 0.2 2.0 • 0.1 2.0 • 0.1 1.3 • 0.1
ATCC 21799(pGC42) c
No induction 0.9 • 0.1 5.6 • 0.3 6.7 • 0.3 1.3 • 0.1 1.0 • 0.1
1.5 mmol of IPTG 0.9 • 0.1 11.8 • 0.6 ( 0.1 4.6 • 0.2 1.9 • 0.2
a From Col6n et al., 1995a,b.
b E. coli-C, glutamicum shuttle vector; pJD4, Km a homar-thrB operon; pGC42, Km a Ap a laqI q
hom ar tac-thrB.
c ATCC 21799(pGC42 ) induced by given amount of IPTG.
6 Examples of Pathway Manipulations 219
the addition of IPTG (Col6n et al., 1993; Col6n et al., 1995a,b). By
increasing the activity of homoserine kinase relative to that of homoserine
dehydrogenase, homoserine secretion essentially was eliminated and the final
threonine titer was increased by about 120% (Table 6.4).
As indicated in Table 6.4, a significant fraction of threonine is either
converted to isoleucine or further degraded to glycine. The conversion of
threonine to isoleucine was prevented by the construction of defined ilvA
mutants via marker exchange mutagenesis (Col6n et al., 1997). At this point
emphasis is placed on blocking the degradation of threonine to glycine. This
is a more challenging problem that involves more than one pathway, the
genes of which have not yet been characterized in Corynebacteria.
Isoleucine
The biosynthesis of isoleucine in C. glutamicum starts with the conversion
of L-threonine to a-ketobutyrate by L-threonine deaminase (LTD, ilvA),
followed by the condensation of this molecule with ce-acetolactate catalyzed
by acetohydroxy acid synthase (AHAS, ilvB-ilvN). This pathway also pro-
vides the precursors for the synthesis of the other two branched-chain amino
acids (BCAA), namely, valine and leucine. Even though up to five different
AHAS isozymes have been reported in enterobacteria, only one enzyme is
known in C. glutamicum. Both LTD and AHAS are inhibited by isoleucine,
whereas AHAS is also inhibited by leucine and valine. Furthermore, all three
BCAA repress the expression of AHAS. The identification and characteriza-
tion of genes involved in BCAA biosynthesis in Corynebacteria have been the
subject of intensive investigation in the last few years (Col6n et al., 1995;
Cordes et al., 1992; Keilhauer et al., 1993).
Overproduction of isoleucine was achieved through the amplification of
the ilvA gene (plasmid pGC77, Table 6.5), in combination with the horn dr
and thrB genes of plasmid pGC42 (see the threonine case). The Corynebac-
terium ilvA gene encoding threonine dehydratase was isolated from a pM2-
TABLE 6.5 Isoleucine Production by C. glutamicum Recombinant Strains a
Excreted amino acids (g L -I)
C. glutamicum strain Lysine Threonine Homoserine Glycine Isoleucine
ATCC 21799(pM2) 22.0 + 1.0 < 0.1 < 0.1 < 0.1
ATCC 21799(pGC42) 0.9 + 0.1 11.8 + 0.6 < 0.1 4.6 + 0.2
ATCC 21799(pGC77) b 0.4 + 0.1 < 0.1 < 0.1 0.5 + 0.1
, .
a From Col6n et al., 1995.
b Derived from pGC42 (Table 6.4) Km RAp R laqI q hom dr ilvA tac-thrB.
< 0 . 1
1.9 + 0.2
15.1 _ 0.2
2 2 0 Metabolic Engineering
based genomic C. glutamicum library by heterologous complementation of an
E. coli ilvA mutant. The resulting plasmid (pGC77), when inserted in the
lysine strain (ATCC 21799), resulted in about 15g L -1 isoleucine, along with
small amounts of lysine and glycine. A carbon balance indicates that the
majority of carbon previously converted to threonine, lysine, glycine, and
isoleucine (21799/pGC42) was incorporated into isoleucine by the new
strain (21799/pGC77).
6.1.3. SOLVENTS
The history of acetone, butanol, and ethanol (ABE) industrial fermentation
processes dates back to the beginning of the century. Due to the shortage of
natural rubber, the English firm Strange and Graham investigated the possi-
bility of manufacturing synthetic rubber. It was then determined that the
synthetic rubber precursors butadiene and isoprene could be best produced
from butanol or isoamyl alcohol. It was in this situation that Professor
Perkins and his assistant Chaim Weizmann (later to become the first presi-
dent of Israel) were recruited to study the chemical production of rubber
precursors. Despite his chemistry education, Weizmann soon concluded that
the key to the success of such a processwas through fermentation; thus, he
retrained himself as a microbiologist. Between 1912 and 1914 he screened
several bacterial strains and succeeded in isolating one, initially termed BY,
that was later termed Clostridium acetobutylicum, which gave the highest
yields of acetone and butanol from starch.
The subsequent development of ABE fermentation processes was acceler-
ated rapidly by the outbreak of World War I, due to the demand for acetone
as the colloidal solvent for nitrocellulose. World War II resulted in a further
demand for acetone and the substrate was changed from maize to molasses,
which was relatively inexpensive and abundant in the 1930s. After World
War II, ABE fermentation declined and virtually ceased in the United States
and United Kingdom with the advent of solvent production from petroleum
and an escalation in molasses prices.
The demise of ABE fermentation was due to a number of intrinsic system
limitations, such as low final concentrations, yields, and productivities,
undesirable solvent ratios, and relatively high substrate costs. Genetic engi-
neering of microbial solvent-producing strains can potentially revive ABE
fermentation processes by addressing the following challenges:
�9 increase product yields and introduce alternative substrates derived
from lignocellulose- or waste-based feedstocks
�9 develop of a strain that exhibits high productivities in continuous and
immobilized cell systems
6 Examples of Pathway Manipulations 221
�9 develop of a strain that gives higher final product concentrations and
exhibits enhanced endproduct tolerance
�9 develop a strain that will easily allow the manipulation of solvent ratios
The induction of several solventogenic enzymes at the onset of solvent
formation suggests that the genetic control of solvent formation is important
(Bennett and Rudolph, 1995; Sauer and Duerre, 1995). However, despite the
cloning and sequencing of several acid- and solvent-associated genes, the
understanding of metabolic flux regulation still remains elusive. Genetic
tools, such as plasmid vectors, have been developed for a number of
clostridial strains. These include (1) cloning vectors that utilize the broad-
host-range conjugal pAM/31 replicon of Enterococcus faecalis or the pIM13
replicon of Bacillus subtilis; (2) suitable selection markers are erythromycin
and clarinthromycin, which is stable at the low prevailing pH; (3) the highly
expressed Clostridium ferredoxin promoter has been exploited in the con-
struction of an expression vector; (4) the conjugative transposons Tn916,
Tn925, and Tn1545 function in clostridial strains, so that transposon muta-
genesis may be possible. Numerous clostridial genes have been cloned and
studied in E. coli (Bennett and Rudolph, 1995; Durre et al., 1995; Papout-
sakis and Bennett, 1991), waiting to be used in gene-inactivation physiologi-
cal studies in Clostridium strains.
The first successful cloning of acid and solvent formation genes in C.
acetobutylicum was reported in 1992 for heterologous overexpression of
acetoacetate decarboxylase (adc) and phosphotransbutyrylase (ptb) in strain
ATTC 824 (Mermelstein et al., 1992) (see Fig. 2.8). This was possible with
the development of a B. subtilis/C, acetobutylicum shuttle vector (pFNK1), in
conjunction with an improved electrotransformation protocol. Acetoacetate
decarboxylase (AADC) is the terminal enzyme in the pathway for acetone
production, converting acetoacetate to acetone and CO 2. Phosphotrans-
butyrylase (PTB) is the branch point enzyme for butyrate production, con-
verting butyryl-CoA and inorganic phosphate into butyryl phosphate
(subsequently converted to butyrate by butyrate kinase) and reduced CoA.
Alternatively, butyryl-CoA is converted into butanol in two enzymatic steps.
AADC activity in the recombinant strain increased by over 9-fold in the
exponential phase and over 33-fold in the stationary phase, whereas PTB
activities of the engineered strain increased by over 20-fold in the exponen-
tial phase and 40-fold in the stationary phase. The transformed strain showed
an increase of 95, 37, and 90% in the levels of acetone, butanol, and ethanol,
respectively. Furthermore, acid concentrations at the end of the fermentation
were considerably lower in the engineered strain (22-fold) than in the
control, and the solvent yield from glucose increased by about 50% in the
redesigned strain.
222 Metabolic Engineering
In a different study, the feasibility of genetic manipulation in Clostridia
was demonstrated by altering the substrate range of C. acetobutylicum
NCIMB 8052: An artificial operon containing the celC and celA genes from
C. thermocellum was transferred to the NCIMB 8052 strain. The resulting
transformant was able to grow on lichenan (a /3-glycan) as the sole carbon
source.
1,3-Propanediol
1,3-Propanediol (1,3-PD) is an intermediate in chemical and polymer
synthesis, e.g., in the synthesis of polyurethanes and polyesters. It is cur-
rently derived from petroleum and is very expensive to produce relative to
similar diols. Tong et al. (1991) recently constructed an E. coli propanediol-
producing strain carrying genes from the Klebsiella pneumoniae dha regulon.
The dha regulon in Klebsiella pneumoniae enables the organism to grow
anaerobically on glycerol and produce 1,3-PD. Escherichia coli, which does
not have a dha system, is unable to grow anaerobically on glycerol without
an exogeneous electron acceptor and does not produce 1,3-PD. In the first
step (see Fig. 6.6), glycerol is converted into 3-hydroxypropionaldehyde by a
~.~_ Glycerol j ) "~__ G lycerOl(dhaB)Dehydratase_~ - I 3-Hyd roxypropionaldehyde .... 1
............................... 1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . I ~ NADH
NAD*
Glycerol
Dehydrogenase
(dhaO)
Dihydroxyaceton
(DHA)
ATP
ADP
i ,
NADH
NAD §
1,3-Propanediol
Oxidoreductase (dha 7")
1
FIGURE 6.6 Pathways for the dissimilation of glycerol in K. pneumoniae. Cloning of K.
pneumoniae dhaB and dhaT genes in E. coli yielded a recombinant strain that converts glycerol
into the industrially useful product 1,3-propanediol.
6 Examples of Pathway Manipulations 223
coenzyme B12 dependent dehydratase, which is then reduced to 1,3-propan-
ediol by an NAD dependent oxidoreductase.
A genomic library of K. pneumoniae ATCC 25955 constructed in E. coli
AG1 was enriched for the ability to grow anaerobically on glycerol and
dihydroxyacetone and was screened for the production of 1,3-PD. The
cosmid pTC1 was isolated from a 1,3-PD-producing strain of E. coli and
found to possess enzymatic activities associated with four genes of the dha
regulon: glycerol dehydratase (dhaB), 1,3-PD oxidoreductase (dhaT), glyc-
erol dehydrogenase (dhaD), and dihydroxyacetone kinase (dhaK) (see
Fig. 6.6). All four activities were inducible by the presence of glycerol. When
E. coli AG 1/pTC1 was grown on complex medium plus glycerol, the yield of
1,3-PD from glycerol was 0.46 mol mo1-1. The major fermentation byprod-
ucts were formate, acetate, and D-lactate. The 1,3-PD fermentation provides
a useful model system for studying the interaction of a biochemical pathway
in a foreign host and for developing strategies for metabolic pathway
engineering.
Further progress in this area is needed in order to minimize byproduct
formation, eliminate the need for glycerol supplementation, and also extend
the substrate range of the pathway to more abundant renewable compounds.
An analysis of the maximum theoretical yield of 0.875 mol 1,3-PD per mol
from glycerol indicates that the yield can be improved. No microorganism is
available that can convert glycerol entirely to 1,3-PD and CO 2 due to theneed for the regeneration of reducing power. Cells regenerate reducing
power (NADH) by forming a mixture of byproducts, such as acetate and
formate, that in essence reduces the maximal theoretical yield to 0.667 mol
mo1-1. In principle, it is possible to provide an alternative source of reducing
power by supplementing glycerol fermentations with pentoses or hexoses.
Theoretical yields of such processes are summarized (moles of 1,3-PD per
mole of glycerol) (Tong and Cameron, 1992):
-2glycerol- glucose + 2 1,3-PD + 2acetate
+ 2formate = 0 (theoretical yield - 1.0 mol mo1-1 )
-5glycerol- 3xylose + 5 1,3-PD + 5acetate
+ 5formate - 0 (theoretical yield - 1.0 mol mo1-1 )
Pilot runs with the E. coli strain carrying the K. pneumoniae dha regulon
indeed resulted in enhanced yields of 1,3-PD from glycerol with cosubstrate
feed: The yield was improved from 0.46 mol mo1-1 with glycerol alone to
0.63 mol tool -1 with glycerol + glucose and 0.55 mol tool -1 with glycerol +
xylose. Such improvements are important economically as the prices of
glucose and xylose are significantly lower than that of glycerol.
2 24 Metabolic Engineering
6.2. E X T E N S I O N OF S U B S T R A T E R A N G E
Most of the work in this area focused on engineering organisms to use
xylose, the primary five-carbon sugar in hemicellulosic biomass, and lactose,
a major byproduct of the dairy industry. Other efforts have examined the
utilization of other plentiful carbon sources, such as whey, starch, and
cellulose. In general, expansion of the ability of microbial strains to utilize a
spectrum of carbonenergy sources provides increased flexibility in the design
and improves the economic feasibility of fermentation processes. This is
particularly true for large-volume commodity operations in which the cost of
the substrate may contribute a very large fraction of the total production cost
(60-65% for ethanol, 40-45% for lysine, and 25-35% for antibiotics and
industrial enzymes). As most microorganisms share a large number of
common metabolic pathways, extension of the substrate range usually in-
volves the addition of only a few enzymatic steps. Occasionally, however,
such steps need to be coordinated with downstream reactions, and it is in
these cases where the tools of metabolic engineering are very useful indeed.
6.2.1. METABOLIC ENGINEERING OF PENTOSE
METABOLISM FOR ETHANOL PRODUCTION
Along with the introduction of ethanol genes in enteric bacteria, parallel
efforts were also undertaken to incorporate pentose-metabolizing pathways
in natural ethanol producers such as S. cerevisiae and Z. mobilis. Microorgan-
isms, in general, metabolize xylose to xylulose through two separate routes
(Fig. 6.7). The one-step pathway catalyzed by xylose isomerase is typical in
bacteria, whereas the two-step reaction involving xylose reductase and xylitol
dehydrogenase is usually found in yeast. Xylulose is subsequently phospho-
rylated by ATP and catabolized via the pentose phosphate pathway and the
EMP pathway (or the ED pathway in organisms such as Z. mobilis). During
the last few years, the genes encoding the enzymes of xylose utilization have
been cloned and characterized in E. coli and some other bacteria (Lawlis
et al., 1984; Rygus et al., 1991). Efforts to isolate natural ethanologenic
microbes that can utilize xylose have not been successful. These provided the
impetus for introducing xylose utilization genes into organisms, especially
those used for ethanol production, and, as such, had the advantage of high
ethanol tolerance.
6 Examples of Pathway Manipulations 225
X . Xylose isomerase J . . . . I Xylulose kinase ,..J - ylose .-, xylulose I ....... Xylulose-5-P
I I
NADDPH" ~ J ~ / ~ NADH Pentose Phosphatepathway
x,,,o., x,,,,o, f
NADP Xylitol NAD Pathway
NAD ~....
ETHANOL
FIGURE 6.7 Xylose metabolism in bacteria and yeast.
Yeast
Even though certain types of yeast such as Pachysolen tannophilus, Pichia
stipitis, or Candida shehatae are xylose-fermenting, they have poor ethanol
yields and low ethanol tolerance compared with the common glucose-
fermenting yeasts, such as S. cerevisiae. Early attempts to introduce the one-
step pathway by cloning the xylose isomerase gene from either E. coli (Sarthy
et al., 1987) or B. subtilis (Hollenberg and Sahm, 1988) in S. cerevisiae were
unsuccessful due to the inactivity of the heterologous protein in the recombi-
nant host cell.
In most yeasts and fungi, xylose reductase and xylitol dehydrogenase are
dependent on NADPH and NAD, respectively (Fig. 6.8). However, examples
of yeast xylose reductases exist that have dual coenzyme specificity (i.e.,
NADPH and NADH), such as those from P. stipitis and C. shehatae. Such a
type of enzyme has the advantage of preventing imbalances of the
NAD/NADH redox system, especially under oxygen-limiting conditions.
Recently, the P. stipitis genes for xylose reductase and xylitol dehydrogenase
were introduced in S. cerevisiae (strain H) (K/Stter et al., 1990; Tantirungkij
et al., 1993). Whereas P. stipitis converts xylose primarily to ethanol under
anaerobic conditions, ethanol production in the recombinant S. cerevisiae
strain (strain H) was marginal (2.7 g L-l), accompanied by the accumulation
of considerable amounts of xylitol (35 g L-l). The observation that ethanol
yield and productivity were higher in aerobic conditions was explained on
the basis of improved NAD regeneration from NADH, which in turn stimu-
lates xylitol dehydrogenase. Additional limitations of xylose utilization in
S. cerevisiae were also attributed to the inefficient capacity of the nonoxida-
tive PPP, as indicated by the accumulation of sedoheptulose-7-P.
226 Metabolic Engineering
12 Xylose XR-
6 NADH ~ X!
6 NAD !
6 NAD ~ I X H
6 NADH
6 Xylulose
. . . .
6 ATP ~ XK
6 ADP 3 F6P
6 X y l u l ~
_1 6Xylitol [
3 Xylulose-5-P
9 ETHANOL
FIGURE 6.8 Anaerobic xylose utilization and cofactor regeneration in recombinant S. cere-
visiae. Abbreviations: EMP, Embden-Meyerhof-Parnas; PPP, pentose phosphate pathway; XR,
xylose reductase; XDH, xylitol dehydrogenase; XK, xylulose kinase.
6 Examples of Pathway Manipulations 227
Further improvement of strain H was attempted via random mutagenesis
and selection for ceils that grow rapidly on xylose (Tantirungkij et al., 1994).
Interestingly, strain IM2, which grew 3 times faster in xylose medium than
strain H, showed lower specific activities of both xylose reductase and xylitol
dehydrogenase, but 1.5 times higher specific activity of xylulose kinase.
Despite the higher growth rate, however, ethanol production by strain IM2
was improved marginally to about 4.2 g L -1 at a yield of 0.08 g g-1.
Zymomonas mobilis
Xylose also could be a useful carbon source for the ethanol producer Z.
mobilis. This is a bacterium that has been used as a natural fermentative
agent in alcoholic beverage production and has been shown to have ethanol
productivity superior to that of yeast. Overall, it demonstrates many of the
desirable traits sought in an ideal biocatalyst for ethanol, such as high
ethanol yield, selectivity and specific productivity, as well as low pH and
high ethanol tolerance. In glucose medium, Z. mobilis can achieve ethanol
levels of at least 12% (w/v) at yields of up to 97% of the theoretical value.
When compared to yeast, Z. mobilis exhibits 5-10% higher yields and up to
5-fold greater volumetric productivities. The notably high yield of this
microbe is attributed to reduced biomass formation during fermentation,
apparently limited by ATP availability. Note that this organism produces only
1 mol of ATP per mole of glucose through the ED pathway [see reaction (2.6)
and Box 6.1] compared with 2 mol for yeast (EMP pathway). As a matter offact, Zymomonas is the only genus identified to date that exclusively utilizes
the Entner-Doudoroff pathway anaerobically. Furthermore, glucose can read-
ily cross the cell membrane of this organism by facilitated diffusion, effi-
ciently be converted to ethanol by an overactive pyruvate decarboxylase/al-
cohol dehydrogenase system, and is generally recognized as a safe (GRAS)
organism for use as an animal feed. As discussed earlier, the main drawback
of this microorganism is that it can only utilize glucose, fructose, and sucrose
and thus is unable to ferment the widely available pentose sugars.
This led Zhang et al. at the National Renewable Energy Laboratory
(Golden, CO) to attempt to introduce a pathway for pentose metabolism in
Z. mobilis (Zhang et al., 1995). Early attempts by other groups using the
xylose isomerase (xy/A) and xylulokinase (xylB) genes (Fig. 6.7) from either
Klebsiella or Xanthomonas were met with limited success, despite the func-
tional expression of these genes in Z. mobilis. It soon became evident that
such failures were due to the absence of detectable transketolase a.ld
transaldolase activities in Z. mobilis, which are necessary to complete a
228 Metabolic Engineering
BOX 6.1
Theoretical Ethanol Yield on Xylose by Recombinant Zymomonas
Strain
The stoichiometry of ethanol production in this recombinant organ-
ism (Fig. 6.8) can be summarized as follows (neglecting the NAD(P)H
balances):
3xylose + 3ADP + 3P i ~ 5ethanol + 5CO 2 + 3ATP + 3H20
Thus, the theoretical yield on ethanol is 0.51 g of ethanol/g of xylose
(1.67 mol mol-1). It is important to note that the metabolically engi-
neered pathway yields only 1 mol of ATP from 1 mol of xylose,
compared with 5 / 3 mol typically produced through a combination of
the pentose phosphate and EMP pathways. The energy limitation
results in less biomass formation and thus a more efficient conversion
of substrate to product.
functional pentose metabolic pathway (Fig. 6.9). After the transketolase
E. coli gene was cloned and introduced in Z. mobilis, a small conversion of
xylose to CO 2 and ethanol occurred (Feldmann et al., 1992). The next step
was to introduce the transaldolase reaction, as this strain accumulated
significant amounts of sedoheptulose-7-P intracellularly. Sophisticated
cloning techniques therefore were applied for the construction of a chimeric
shuttle vector (pZB5) that carries two independent operons: the first encod-
ing the E. coli xylA and xylB genes and the second expressing transketolase
(tktA) and transaldolase (tal) again from E. coli. The two operons compris-
ing the four xylose assimilation and nonoxidative pentose phosphate path-
way genes were expressed successfully in Z. mobilis CP4. The recombinant
strain was capable of fast growth on xylose as the sole carbon source, and
moreover it efficiently converted glucose and xylose to ethanol with 86 and
94% of the theoretical yield from xylose and glucose respectively. This
represents a complementary approach to the previously discussed expression
of the Z. mobilis PET operon in E. coli for ethanol production.
6 Examples of Pathway Manipulations 2 2 9
I / Xylulose < XDH [ Xylitol p XR - Xylose
Footo .o. j
XK . . . .
TK
.1 �9 ] Xylulose-5-P Sedoheptulose-7-P Erythrose--P
l 1
t
TK
"J Glyceraldehyde-3-P ! "!
Pyn rate
[Acetaldehyde I
ETHANOL
FIGURE 6.9 Ethanol production from pentose sugars in metabolically engineered Z. mobilis.
Appreviations: XR, xylose reductase; XDH, xylulose dehydrogenase; TK, transketolase; TA,
transaldolase.
6 . 2 . 2 . CELLULOSE- HEMICELLULOSE
DEPOLYMERIZATION
It would be desirable if ethanol-producing microbes from lignocellulose also
had means to depolymerize cellulose, hemicellulose, and associated carbohy-
drates. Many plant pathogenic bacteria (soft-rot bacteria), such as Erwinia
carotovora and Erwinia chrysanthemi, have evolved sophisticated systems of
hydrolases and lyases that aid the solubilization of lignocellulose and allow
230 Metabolic Engineering
them to macerate and penetrate plant tissue (Kado, 1992). Genetic engineer-
ing of these bacteria for ethanol production represents an attractive alterna-
tive to the solubilization of lignocellulosic biomass by chemical or enzymatic
means. E. carotovora SR38 and E. chrysanthemi EC16 were genetically
engineered with the PET operon and shown to produce ethanol and CO 2
efficiently as primary fermentation products from cellobiose, glucose, and
xylose (Beall and Ingram, 1993). Both ethanologenic Erwinia strains pro-
duced about 50 g L -1 ethanol from 100 g L -1 cellobiose in less than 48 hour
with a maximum volumetric productivity of 1.5 g of ethanol L -1 hour. This
rate is over twice that reported for the cellobiose-utilizing yeast, Bret-
tanomyces custersii, in batch culture (Spindler et al., 1992).
Along similar lines, the incorporation of saccharifying traits to ethanol-
producing microorganism was also attempted. The gene encoding for the
xylanase enzyme (xynZ) from the thermophilic bacterium C. thermocellum
was expressed at high cytoplasmic levels in ethanologenic strains of E. coli
KOll and K. oxytoca M5A1(pLOI555) (Burchhardt and Ingram, 1992). This
is a temperature stable enzyme that depolymerizes xylan to its primary
monomer (99%) xylose. In order to increase the amount of xylanase in the
medium and facilitate xylan hydrolysis, a two-stage, cyclical process was
employed for the fermentation of polymeric feedstocks to ethanol by a single,
genetically engineered microorganism. Cells containing xylanase were har-
vested and added to a xylan solution at 60~ thereby lysing and releasing
xylanase for saccharification. After cooling to 30~ the hydrolysate was
fermented to ethanol, in the meantime replenishing the supply of xylanase
for the subsequent saccharification. K. oxytoca was found to be a superior
strain for such an application, because, in addition to xylose (metabolizable
by E. coli), it can also consume xylobiose and xylotriose. Even though the
maximum theoretical yield of M5A1(pLOI555) is in excess of 48 g L -1
ethanol from 100 g L -1 xylose, about one-third of that was achieved in this
process because xylotetrose and longer oligomers remained unmetabolized
by this strain. The yield appeared to be limited by the digestibility of
commercial xylan rather than by the lack of sufficient xylanase activity or by
ethanol toxicity.
6 . 2 . 3 . LACTOSE AND WHEY UTILIZATION
Whey is a nutrient-rich byproduct of the dairy industry that can provide an
inexpensive carbon and nitrogen source in biotechnological processes. It
reaches an annual production of 1011 kg, with high lactose (75% of dry
matter) and protein contents (12-14%), as well as small amounts of organic
acids, minerals, and vitamins. Whereas its protein content is separated and
6 Examples of Pathway Manipulations 231
~ 1 ~ 8 0 - 9 0 % (w/w)
WHEY
(>115 million tons / year)
Lactose: 4-5%
Protein: 0.7%
Fat: 0.3%
Mineral Salts: 0.4-0.5%
Soluble Vitamins
V Ih,.
A v
Waste Disposal
High Chemical Oxygen Demand
(COD: 60-80 g liter 1)
• Wheyi I Enzyme i >; Hydrolysis ; " ~ c o ~ " ....... 7Proteins] >I Peptides J-----
Enzyme
J J ~. Hydrolysis Permeate I (~-Galactosidase)
r
robic
ntation
r
tool
:erol
,'one
tool
Various
Fermentations
. . . . .
Aminoacids
Lactate, Acetate
Citric Acid
Xanthan gum
I Glucose I i
Galactose Syrup
.......
J Food Applications
~ Aerobic
ermentation
~ , ,
Biom~,s
Enzymes
Lipids
Pigments
. . . . .
Anaerobic
Digestion
1
t - ~ 1 7 6 , I
t Anae
Ferme
Eth~
Gly(
Ace'
Butl
FIGURE 6.10 Utilization of whey components in fermentation processes (vonStockar and
Marison, 1990).
concentrated for food purposes (see Fig. 6.10), the lactose and salts in the
permeate have lower values and are typically discarded. In addition to the
loss of valuable nutrients, disposal also requires expensive sewage treatment.
Efforts therefore are intensifying to find useful applications for whey in
general and for the permeate in particular. Some examples of fermentation
processes that can utilize whey byproducts as feedstocks are summarized in
Fig. 6.10.
Although a variety of microbes can utilize whey, some of the industrially
most prominent organisms such as S. cerevisiae, Z. mobilis, and Alcaligenes
eutrophus are unable to do so. Utilization of lactose requires the presence of
the catabolic enzyme ~8-galactosidase (coded by the lacZ gene) for the
hydrolysis of the lactose disaccharide into its constituent sugars, glucose and
galactose. Additionally, an efficient lactose transport system, along with
232 Metabolic Engineering
glucose- and galactose- catabolizing pathways, is needed. These requirements
were evident in the early work of introducing the lactose transposon, Tn 951
from Yersinia enterocolitica (harbors lacI, lacZ and lacY genes: see Chapter 5
section on the lac operon), in Z mobilis (Carey et al., 1983; Goodman et al.,
1984). Although the E. coli ]3mgalactosidase was successfully expressed in
this strain, ethanol yields were much lower than theoretical values for at least
two reasons: lactose is cleaved by ]3-galactosidase to glucose and galactose;
however, only glucose can be fermented to ethanol by the recombinant
Z. mobilis strains, with galactose accumulating to inhibitory concentrations
(Yanase et al., 1988). This indicates that the galactose operon is needed in
addition to the lactose operon. Secondly, the poor ethanol productivity was
attributed to the slow uptake of lactose. Later studies utilized the Tn 951 to
express ~8-galactosidase in Pseudomonas saccharophila and Alcaligenes eutro-
phus (Pries et al., 1990). This allowed transconjugants of P. saccharophila to
grow slowly on lactose mineral medium whereas the parent strain did not
grow at all. Plasmid pPL76, harboring an A. eutrophus promoter-lacZ fusion,
enabled A. eutrophus not only to express jS-galactosidase but also to grow
slowly on lactose. Subsequently, the E. coli gal operon was also transferred
to these strains to allow galactose utilization.
The E. coli lacZY operon (coding for jS-galactosidase and lactose perme-
ase) was also integrated into the Pseudomonas aeruginosa chromosome, an
important producer of rhamnolipid biosurfactants (Koch et al., 1988). The
transconjugants grew well in lactose-based media (minimal medium and
whey) and produced rhamnolipid during the stationary phase. The E. coli
lacZY gene, under the control of phage ~LO promoter, was also inserted in
Xanthomonas campestris, a bacterium that causes tremendous agricultural
losses worldwide but also used in the production of xanthan gum (Fu and
Tseng, 1990). For the production of xanthan, glucose, sucrose, or starch
media are normally used. The recombinant strain, however, expressed high
levels of ~8-galactosidase and grew well in a medium containing lactose as the
sole carbon source. Production of xanthan gum in lactose or diluted whey by
the engineered strain was evaluated, and it was found to produce as much
xanthan gum using these substrates as did cells in a glucose medium. These
examples illustrate attractive processes for treating industrial waste materials
while producing useful compounds at the same time.
An alternative strategy is to construct a strain that secretes ~8-galactosidase
into the medium or into the periplasm wherein lactose is freely diffusible.
This approach was applied in yeast, where a lactose utilizing S. cerevisiae
was constructed by expressing the gene for a secreted, thermostable ~8-galac-
tosidase (lacA) from Aspergilus niger (Kumar et al., 1992). This study
demonstrated that 40% of the total recombinant protein was secreted into
the medium, allowing S. cerevisiae to grow on whey permeate (4% w / v
lactose) with a doubling time of 1.6 h. This approach offers significant
6 Examples of Pathway Manipulations 233
advantages over the earlier processes for the fermentation of whey by S.
cerevisiae, which used either ]3-galactosidase prehydrolyzed whey or yeast
co-immobilized with ]3-galactosidase.
The entire E. coli lactose operon was also inserted into the amino acid
producer C. glutamicum R163 (Brabetz et al., 1991). Recombinant C. glutam-
icum strains carrying their lac genes under the control of a strong promo-
ter grew rapidly in defined media with lactose as the sole carbon source
(3% w/v- l ) . The growth characteristics, which were indistinguishable from
those in glucose media, depended on the presence of the lacY gene (lactose
permease) in addition to lacZ. Furthermore, enzymatic assays indicated that
all ]3-galactosidase activity was intracellular. Again, the main drawback of
this system is the inability of the cells to utilize the second monosaccharide
of lactose, namely, galactose.
6 . 2 . 4 . SUCROSE UTILIZATION
Sucrose (a disaccharide of glucose and fructose) is another abundant and
inexpensive carbon source found in cane molasses, for example. Even though
certain E. coli strains can utilize sucrose, E. coli K-12, a potentially useful
industrial organism for amino acid production, is unable to grow on sucrose.
Various researchers have attempted to express the sucrose utilization system
(Scr +) from other bacterial species, but are unable to stably maintain the
Scr + phenotype in E. coli. Recently, a successful attempt came about from
the cloning of the scrA gene coding for sucrase from E. coli B-62 onto a
plasmid and then transferring the cloned DNA fragment onto the chromo-
some of E. coli K-12. Tryptophan producer derivatives of E. coli K-12
expressing the scrA gene grew well in sucrose medium and excreted amounts
of tryptophan (5.7 g L -I) comparable to these from similar strains grown on
glucose (Tsunekawa et al., 1992).
6.2.5. STARCH = DEGRADING MICROORGANISMS
Starch, derived from renewable resources such as corn and cereals, is a very
important carbon and energy source in biotechnological processes. Substitu-
tion of glucose with starch not only can reduce fermentation feedstock costs
but also can minimize or eliminate negative physiological effects associated
with glucose, such as catabolic repression or acidogenesis.
Starch is a mixture of linear and branched homopolymers of D-glucose
that are connected by a(1 ~ 4) linkages and at branch points by a(1 ~ 6)
linkages. It is formed as a carbohydrate reserve in plants and is present in
234 Metabolic Engineering
significant amounts in potato tubers and in the seeds of wheat, corn, barley,
and sorghum. The linear component, amylose, consists of chains of c~-I,4-D-
glucopyranose ranging in degree of polymerization from about 102 to 4 x
10 ~ In the branched component, amylopectin, shorter chains (17-23 units
long) of c~-l,4-D-glucopyranose are linked together by a-1,6 bonds to form a
branched structure with a degree of polymerization ranging from 10 4 tO 4 x
107. Four types of starch-decomposing enzymes are of importance: c~-amylase,
]3-amylase, pullulanase or isoamylase (debranching enzymes), and glucoamy-
lase. a-Amylase is an endoglucanase that randomly cleaves c~(1 -~ 4) link-
ages, converting starch to dextrins, maltose, and glucose. It is produced by
bacteria and fungi, notably by Bacillus species, Pseudomonas, and Lactobacilli
and by Aspergillus species. ~-Amylase is an exoglucanase typically found in
plants that successively removes maltose units from the nonreducing end of
starch. Pullulanase and isoamylase belong to the debranching group of
enzymes thathydrolyze c~(1 -~ 6) linkages. Glucoamylase is a fungal enzyme
that removes glucose residues from the nonreducing end of starch.
Because most microorganisms are unable to degrade this glucose biopoly-
mer, work has focused on cloning genes for enzymatic starch hydrolysis into
various organisms (Kennedy et al., 1988). This approach offers an attractive
alternative to current processes that first convert starch enzymatically into
glucose and some oligosaccharides and then use them as carbon sources in a
separate fermentation step. Along these lines, a S. cerevisiae strain was
constructed that contained a glucoamylase gene from Aspergillus sp. (Innis
et al., 1985). The recombinant strain was able to grow on amylodextrins,
albeit at a lower rate than the case where glucoamylases are added added to
the fermentation medium.
Useful applications of such a recombinant strain include brewing and
baking. In the case of brewing, the malting process, the partial hydrolysis of
barley starch, results in a considerable amount of dextrins that cannot be
fermented by the yeast S. cerevisiae. These dextrins are of high caloric
content and have to be removed for the production of light beer, presently
achieved by the external addition of glucoamylase. Therefore, an engineered
strain with amylolytic properties would offer a suitable alternative for
brewing, especially for the production of a low-calorie product. Also, such a
strain would eliminate the need for c~-amylase-enriched flour in certain types
of bread manufacturing.
Strains with the proceding desirable characteristics recently were con-
structed by expressing the yeast Schwanniomyces occidentalis c~-amylase
(AMY1) and glucoamylase (GAM1) genes in S. cerevisiae (Hollenberg and
Strasser, 1990). Comparative enzymatic studies illustrated that the engi-
neered amylolytic system is as effective as the original S. occidentalis strain.
During fermentation of ground, liquefied wheat, this recombinant strain
6 Examples of Pathway Manipulations 235
showed the same ethanol production rate as a conventional distillery yeast
with saccharifying enzymes added prior to fermentation.
6.3. EXTENSION OF PRODUCT SPECTRUM
AND NOVEL PRODUCTS
This is an area with immense potential for metabolic engineering. Rational
expression of heterologous genes can extend existing pathways of the host
organism for the overproduction of both known and novel compounds with
attractive chemical and/or physical properties.
6.3.1. ANTIBIOTICS
Antibiotic production by microorganisms is one of their more interesting
features, particularly from a medical and commercial point of view. More
than 10,000 antibiotics and similar bioactive metabolites have been isolated
from microbes, with approximately 500 new classes of low-molecular-weight
compounds published every year. In monetary terms, antibiotics currently
are the most important group of microbial biotechnological products, with
an estimated world sales in excess of $15 billion. The primary classes are
cephalosporins, penicillins, and tetracyclines, and the majority of these
agents are produced by Streptomyces (and other actinomycetes) and various
Bacillus species. Their primary use is in the treatment of human infectious
diseases, although a significant number have agricultural and veterinary
applications.
Antibiotics are made by secondary metabolic pathways that use com-
mon metabolites in less specific and, sometimes, more intricate ways than
primary metabolism. The polyketide antibiotics, for example, are made from
simple fatty acids by a pathway that superficially resembles the one used to
make long-chain fatty acids, but the resulting compounds exhibit a range of
structural complexity far surpassing the simple hydrocarbon framework
of the essential fatty acids. Recently, it has become apparent that yields of
secondary metabolites, including antibiotics, can also be improved by over-
coming rate-controlling biosynthetic steps through genetic techniques. In
addition, metabolic engineering techniques are applied in order to modify
known antibiotics to improve their properties and also to synthesize new
forms of antibiotics (Summers et al., 1992). For years antibiotic production
in filamentous fungi and Streptomyces was improved by random
mutation/screening and, to a lesser extent, by selecting mutants that over-
236 Metabolic Engineering
produced primary metabolic precursors of antibiotics. Work spanning four
decades developed production strains less amenable to improvement by these
traditional techniques.
The application of recombinant DNA technology was based on the devel-
opment of genetic transformation systems for/3-1actam-producing organisms
and cloning of biosynthetic genes. The ability to transform industrially
important organisms (such as P. chrysogenum and C. acremonium) provided a
powerful tool for the precise manipulation of biosynthetic pathways and an
avenue for practical applications, such as gene dosage studies for possible
limiting steps or gene disruptions to alter final products. The discovery that
many antibiotics genes are clustered, and also that certain genes of related
pathways exhibit cross-hybridization, has opened new avenues in this area
(Charter, 1990). Gene clustering facilitates cloning, and the fact that these
genes are often positively regulated increases the possibility of improving
production through overexpression of the genetic regulatory molecule. Over-
expression of such regulatory genes caused, for instance, the overproduction
of streptomycin, undecylprodigiosin, and actinorhodin in wild-type strains
(Charter, 1990).
Streptomyces rank near the top among microorganisms of industrial
importance, especially as antibiotic producers. Actinorhodin biosynthesis
genes were transferred from Streptomyces coelicitor, the only species with
well-established genetics, to Streptomyces lividans, enabling the latter strain
to produce actinorhodin. Later on, clustered erythromycin genes from Strep-
tomyces erythreus were transferred to S. lividans, allowing the recombinant
strain to produce erythromycin A. Transformation of the fungi Neurospora
crassa and Aspergillus niger with a cosmid containing P enicillium chryso-
genum penicillin biosynthetic genes resulted in the production of penicillin V
by these strains.
Yield improvements through metabolic engineering have been demon-
strated for a number of systems. For example, the production of cephalosporin
C by Cephalosporium acremonium was increased by 15% by overexpressing
the cefEF gene (Skatrud et al., 1989). This gene codes for a bifunctional
protein with two sequentially acting activities: deacetoxycephalosporin C
synthetase and deacetylcephalosporin C synthetase (DACS). The recombi-
nant strain with a 2-fold increase in DACS activity, was able to convert
penicillin N, a precursor normally excreted in large quantities, into the final
product cephalosporin C. This work also identified DAC acetyltransferase,
the final enzyme in the cephalosporin C pathway, as a potentially controlling
step, as a substantial amount of deacetylcephalosporin (DAC) was observed
in the medium.
Recombinant DNA techniques can also be utilized to engineer hybrid or
even novel antibiotics. The main inherent obstacle in such applications is, of
6 Examples of Pathway Manipulations 237
course, the fact that the producing organism must be resistant to the hybrid
antibiotic in order to achieve high yields. Genes for biosynthetic steps in
different organisms can be combined in the same organism, thus extending
the diversity of natural antibiotics. In an early attempt, part of the cloned
pathway for actinorhodin from Streptomyces coelicitor was transformed
into a Streptomyces strain that produces the compound medermycin
(Hopwood etal., 1985). The recombinant strain produced a hybrid antibiotic
identified as mederrhodin. Conversion of the native medermycin to meder-
rhodin involves a ]3-hydroxylation step postulated to be catalyzed by heterol-
ogous ~8-hydroxylation activity of an enzyme with a broad substrate speci-
ficity. McAlpine et al. have used a similar strategy to transform a mutant of
Saccharopolyspora erythraea, which is blocked in an early step of erythro-
mycin biosynthesis, with a DNA library from the oleandromycin producer
Streptomyces antibioticus (McAlpine et al., 1987). One of the recombinant
strains produced an antibiotic with a novel structure, called 2-noreryth-
romycin. A greater challenge in generating novel antibiotics goes beyond
single-group substitutions and involves the alteration of their backbone
structure. Streptomyces galilaeus normally produces aclacinomycin A and B.
Following its transformation with polyketide synthase genes, clones were
obtained that produced anthraquinone (Bartel et al., 1990). These exciting
results provide the foundation for ongoing efforts to rationally design and
synthesize novel antibiotics.
6.3.2. POLYKETIDES
Polyketides are found in most organisms and are especially abundant in a
class of filamentous bacteria, the actinomycetes. The polyketide family is a
rich source of bioactive molecules with antibiotic (such as tetracycline and
erythromycin) and pharmacological (e.g., cancer agents and immunosupres-
sants) properties. Synthesis of these molecules involves giant modular en-
zymes known as polyketide synthases (PKS). Polyketides are made from
simple fatty acids by a pathway that superficially resembles the one used to
make long-chain fatty acids, but the resulting compounds exhibit a range of
structural complexity far surpassing the simple hydrocarbon framework of
biological fatty acids. A major distinction of fatty acid synthesis is the fact
that the initial condensations (/3-keto acid reduction, dehydration, hydro-
genation) do not occur in a regular fashion, but rather depend on the
modular structure of the given polyketide synthase. In fatty acid synthesis,
an acetyl group is added at each round of synthesis to produce a long
unbranched chain, and the carbonyl group introduced at each round is
reduced to CH 2. In the biosynthesis of polyketides, the unit added is often
238 Metabolic Engineering
larger than an acetyl (e.g., malonyl-CoA), yet each condensation step adds
two carbons to the elongating chain in a way that the remaining part of the
unit extends from the main chain as a branch. Some of the carbonyl groups
are not reduced at all, and others are reduced only to the level of CHOH.
Reasons that make polyketides an attractive study model for metabolic
engineering include the following: (1) their complex structure results from
simple units combined in diverse ways; (2) the modular construction of the
enzymatic catalyst (PKS) allows control of enzyme structure and, hence,
polyketide type at the genetic level. Recent progress in this area has estab-
lished the groundwork to generate novel polyketide structures through
genetic engineering of polyketide synthases and, at the same time to derive
knowledge that elucidates the structure-function relationship in polyketide
synthases (Kao, 1997; McDaniel et al., 1993). Moreover, this system provides
an opportunity to bridge the fields of genetics and chemistry and, above all,
promises to enable scientists to rationally design novel molecules at the level
of DNA.
Erythromycin Production by Saccharopolyspora erythrea
The production of this polyketide in S. erythrea involves only three giant
genes, each of which codes for a protein of more than 300 kDa (Cane et al.,
1983, 1987). Each protein is in turn made up of two inexact repeats that can
be divided into six modules, as shown in Fig. 6.11. Each module contains
combinations of at least six monofunctional polypeptides, each responsible
for one single reaction step: acyl transferase (AT), acyl-carrier proteins (ACP),
]3-ketoacyl-ACP-synthase (KS), ]3-ketoacyl-ACP-reductase (KR), dehydrase
(DH), and enoyl reductase (ER) (Fig. 6.11). An interesting and perhaps
anticipated observation is the fact that some of these genes are highly
homologous with genes of the fatty acid biosynthesis pathway. What is
fascinating about this type of organization is the fact that novel molecules
can be generated by using different combinations and permutations of these
basic modules, as well as by introducing point mutations within functional
domains. Nature has already produced a vast diversity of polyketides by this
same technique. A major contribution of metabolic engineering in this area is
to design chemical structures of potentially useful molecules, currently
known or not, by "genetic design."
In the last few years, Khosla and colleagues have focused their attention
on deciphering the rules of polyketide synthetases by developing a Strepto-
myces host-vector system for the expression of recombinant polyketide
synthases (PKS) (Kao, 1997; McDaniel et al., 1993). This work has led to the
concept of a minimal polyketide forming-system containing the condensing
enzyme, the acyl carrier protein, and the malonyl-CoA transferase (McDaniel
6 Examples of Pathway Manipulations 239
Module 1 Module 2
h
! AT AcP KS AT KR ACP KS AT KR ACP "~ I
I v' ! . . . . . . I I I
S S S
I I I
o , i,
I "
CH 3 CHOH CHOH
I I
CH 2 CH-CH 3
I I
CH3 CHOH
I
CH~
I
CH 3
Module 3 _M._oflgJg~ Module 5
KS AT ACP KS'AT DH ER' KR ,cP~l KS AT KR
I I I / I
S S S
kio k !o io q
ICH'CH31 I C H - C H 3 CH'CH3 I
C=0 CH 2 C H O H
I I
CH-CH 3 CH-CH 3 CH-CH 3
i i
CHOH C=0 CH 2
I I
CH-CH 3 CH-CH 3 CH-CH 3
I I
CHOH CHOH C=0
I I
CH 2 CH-CH 3 CH-CH 3
! I
CH 3 CHOH CHOH
I I
CH 2 CH-CH 3
I I
CH 3 CHOH
I
CH 2
I
CH 3
Module 6
S
!
'!, ............. I 21,~176 I
3 CHOH
I
4 OH-OH 3
I
5 CHOH
6 CH'CH 3
7 CH2
8 CH-CH 3
9 C=O
10 CH-CH 3
1 1 CHOH
I
1 2 CH-CH 3
I
13CHOH
I
14CH 2
I
15CH 3
FIGURE 6.11 The organization of genes for erythromycin A biosynthesis in Saccharospora
erythrea. The DNA region is divided into three open reading frames (ORFs), each coding for a
large, complex enzyme molecule. In turn, each enzyme can be subdivided into two modules,
with each successive module adding a new propionic acid unit (box) to the growing chain.
Subunits: acyl transferase (AT), acyl-carrier proteins (ACP), ~-ketoacyl-ACP-synthase (KS),
~8-ketoacyl-ACP-reductase (KR), dehydrase (DH), and enoyl reductase (ER).
et al., 1994). Additional proteins may then function either as chain length
factors, which determine the extent of elongation, or as cyclases, which
direct the mode of cyclization (Hutchinson and Fujii, 1995). A number of
polyketide molecules were produced recently by Streptomyces strains trans-
formed with various combinations of PKS genes comprising minimal sys-
tems, thus paving the way for combinatorial biosynthetic approaches (Shen
and Hutchinson, 1996; Tsoi and Khosla, 1995). Characterization of these
metabolites has provided new insights into the programming aspects of PKS
genes (Box 6.2).
240 Metabolic Engineering
BOX 6.2
Examples of Minimal Polyketide-Forming Systems and Programming
Rules of PKS
S. coelicolor A3(2) is an actinomycete with well-developed genetics
that produces the blue-pigmented polyketide actinorhodin. The act
PKS gene cluster has been cloned and completely sequenced, and, in
addition, a S. coelicolor strain (CH999)was constructed by deleting the
entire act cluster through homologous recombination. This mutant
strain was transformed by plasmid carrying combinatorial "minimal"
gene clusters of various PKSgenes in order to elucidate the mecha-
nisms by which PKSs achieve their high degree of specificity.
For example, the recombinant strain CH999/pRM37 expresses a
"minimal" act PKS gene cluster together with a minimal gene set for
tetracenomycin (tcm) PKS (see figure; AT, acyl transferase; ACP, acyl-
carrier proteins; KS, ]3-ketoacyl-ACP-synthase; KR, ]3-ketoacyl-ACP-
reductase; CYC, cyclase; OMT, o-methyltransferase). The actinorhodin
(act) PKS catalyzes chain termination after nine condensation cycles,
whereas tcm (PKS) does so after nine cycles. This particular strain was
found to produce two novel aromatic polyketides, whose structures
were determined by 1H and 13C NMR. Similar experiments have been
repeated using various minimal gene clusters followed by the identifi-
cation and structural analysis of the resulting polyketide(s).
"minimal" gene cluster for actinorhodin
polyketide synthase (act PKS)
Module 1 Module 2
........ K ~ I I ..... KS AT CLF
Module 3
minimal gene cluster for tetracenomycin
polyketide synthase (tcm PKS)
Module 1 Module 2
, _
6 Examples of Pathway Manipulations 241
Some of the primary conclusions from such studies are summarized
(McDaniel et al., 1993):
�9 The chain length is, at least in part, dictated by a specific protein,
which has been given the name "chain length determining factor"
(CLF)
�9 Some heterologous ketosynthase/putative acyl transferase
(KS/AT) CLF pairs give rise to functional PKSs, but other pairs
are monofunctional.
�9 Acyl-carrier proteins (ACP) could be interchanged among differ-
ent synthases without affecting product structure.
�9 A specific ketoreductase (KR) reduced polyketide chains of dif-
ferent lengths, probably after the complete polyketide chain had
been synthesized.
�9 Regardless of chain length, this ketoreductase reduces the C-9
carbonyl from the carboxyl end.
�9 The regiospecificity of the first cyclization is controlled by the
KS/AT and/or the CLF.
�9 A specific cyclase (CYC), responsible for catalyzing the second
cyclization reaction, evidently can discriminate between interme-
diates of different chain lengths and degrees of reduction.
Such findings form the basis for the systematic exploration of struc-
ture-function relationships in these complex systems and the rational
design of novel polyketides that are based on minimal polyketide-for-
ming systems.
With the further elucidation of PKS strategies, it is envisioned that
combinatorially generated PKS systems will allow the synthesis of polyketide
libraries that contain thousands of new molecules. These libraries could then
be screened for molecules with any type of property, ranging from pharmaco-
logical to materials. Clearly, the upcoming years of modular PKS research
promise to be very exciting ones, especially when one considers the richness
and engineering potential of these fascinating enzyme systems.
6.3.3. VITAMINS
Vitamin C
Established commercial production of the vitamin C (ascorbic acid)
precursor 2-keto-L-gluconic acid (2-KLG) involves a two-stage fermentation
24 2 Metabolic Engineering
Erwinia herbicola Corynebacterium sp.
v r -
CHO C00H COOH COOH COOH
HO~ OH HO~ OH HO~ O HO~ O HO~o'
CH2OH CHzOH CH2OH CH2OH CH2OH
G GA 2-KDG 2,5-KDG 2-KLG
Erwinia herbicola I dkgr
FIGURE 6.12 Biological conversion of glucose (G) to 2-keto-L-gluconic acid (2-KLG). Com-
parison of a two-stage process involving Erwinia herbicola and Corynebacterium sp. with a single
step process based on 17.. herbicola expressing heterologous diketo-D-gluconate reductase (DKGR).
Intermediates: GA, gluconic acid; 2-KDG, 2-keto-D-gluconic acid; 2,5-DKG, 2,5-diketo-D-glu-
conic acid.
process. The first utilizes Erwinia herbicola to convert glucose to 2,5-diketo-
D-gluconate, which is subsequently converted to 2-KLG in a fermentation
step by a Corynebacterium sp. In an effort to change this into a one-stage
process, the E. herbicola was genetically transformed with the Corynebac-
terium gene that encodes 2,5-DKG reductase (DKGR), which catalyes the
2,5-DKG to 2-KLG conversion (Anderson et al., 1985; Grindley et al., 1988)
(Fig. 6.12). After optimizing the culture conditions, these recombinant
strains of Erwinia produced about 120 g L -1 2-KLG within 120 with a molar
yield from glucose of about 60%. Followup studies illustrated the potential
economic advantages of the metabolically engineered strain for vitamin C
production and have led to a number of U.S. patents (Anderson et al., 1991;
Hardy et al., 1990).
Biotin
Biotin is an essential nutrient for many microorganisms and animals. It
acts as cofactor for enzymes involved in fatty acid and carbohydrate
metabolism and is used in animal feed and as an additive in industrial
fermentation processes. Currently, biotin is produced by a complicated and
expensive chemical synthesis method. Even though current economics favor
chemical synthesis, further improvements in microbial biotin production
processes could make bioconversions competitive with existing technologies.
The metabolic pathway for biotin synthesis from pimelic-CoA was first
6 Examples of Pathway Manipulations 243
described in E. coli (Barker and Campbell, 1980), and all enzymes involved
in biotin synthesis from pimelic acid in B. sphaericus were identified (Izumi
et al., 1981). The finding that B. sphaericus secreted significant quantities of
biotin pathway intermediates led to the isolation of the bio genes from this
organism. The genes involved in biotin synthesis, organized in two clusters
bioXWF and bioDAYB, have recently been cloned on E. coli vectors (Gloecker
et al., 1990). E. coli transformed with these genes produced up to 457 mg L -1
of biotin and 350 mg L -1 biotin intermediates (Sabati~ et al., 1991).
Vitamin A
Another example of the application of metabolic engineering to convert
native metabolic intermediates to desirable endproducts is the production of
]3-carotene precursor for vitamin A. In the past, many species of algae and
fungi (e.g., Neurospora crassa, Penicillium sclerotiorum, Phycomyces
blakesleeanus) and also yeasts (Rhodotorula) were considered for use in
]3-carotene production, but were found to be unsuitable (Ninet and Renaut,
1979). Because the precursor for carotenoid biosynthesis, geranylgeranyl
pyrophosphate, exists in many organisms for the synthesis of sterols,
hopanoids, and terpenes, it can be utilized with appropriate genetic engineer-
ing to produce ]3-carotene. Recently, the Erwinia uredovora genes for the
biosynthesis of cyclic carotenoids including /3-carotene have been cloned
and analyzed (Misawa et al., 1990). Following the genetic transformation of
Z. mobilis and Agrobacterium tumefaciens with four of the ]3-carotene genes,
yellow colonies were obtained on agar plates (Misawa et al., 1991). Even
though neither strain is a native producer of ]3-carotene, the transconjugants
produced 220-350 mg DW of the vitamin A precursor in liquid culture. It is
also suggested that ]3-carotene-producing Z. mobilis strains, which is used on
a large scale for ethanol production, can be subsequently used as an animal
feedstock due to its enhanced nutritional value.
In a related effort, again involving heterologous expression of six of the
Erwinia carotenoid genes, an array of geranylgeranyl pyrophosphate byprod-
ucts was obtained in S. cerevisiae. One or more of the following products was
detected, depending on the number of genes in the linear pathway that were
actually expressed: phytoene, lycopene, ]3-carotene, zeaxanthin, and zeaxan-
thin diglucoside (Ausich et al., 1991).
6.3.4. BIOPOLYMERS
Improvement of polymer production by organisms (e.g., xanthan gum and
bacterial cellulose), as well as the production of new biological polymers,is
244 Metabolic Engineering
yet another major application of metabolic engineering (Peoples and Sinskey,
1990). Approximately 93% of fossil resources consumed in the world is for
energy production, while only 7% is used by industries for the production of
a variety of organic chemicals, including solvents and plastics (Eggersdorfer
et al., 1992). Replacement of a fraction of synthetic plastics with biodegrad-
able polymers produced from renewable resources is, thus, likely to have
only a marginal impact on the overall consumption of fossil fuels. Greater
use of biodegradable plastics could, however, significantly contribute to
solving problems associated with environmental pollution and waste man-
agement. The same intrinsic qualities of durability and resistance to degrada-
tion that have made plastics ideal industrial and consumer materials are now
regarded as a source of environmental and waste management problems. In
contrast, biodegradable polymers are either partly or fully composed of
material that can be degraded either by nonenzymatic hydrolysis or by the
action of enzymes secreted by microorganisms. Although some polymers,
such as blends of starch and polyethylene, are only partly biodegradable,
polymers such as poly(3-hydroxybutyric acid) [P(3HB)] are 100% biodegrad-
able as they can be converted into carbon dioxide and energy by microorgan-
isms, such as bacteria, fungi, and algae. More than a dozen biodegradable
plastics are now on the market, representing a range of properties suitable for
various consumer products, with estimates of the current global market for
these biodegradable plastics of up to 1.3 billion kg per year (Lindsay, 1992).
Poly(hydroxyalkanoate)s
Among the various biodegradable plastics available, poly(hydroxyal-
kanoate)s (PHAs) are attracting growing interest. PHAs are a class of intracel-
lular carbon and energy storage materials accumulated by numerous bacteria
in response to environmental limitations (e.g., oxygen or nitrogen depriva-
tion and sulfate or magnesium limitation). Changes in environmental condi-
tions often cause dramatic shifts in intermediary metabolism. Many of these
shifts are controlled by global regulatory networks capable of coordinated
induction or repression of enzyme repertoires (Chapter 5). These polymers
have recently attracted considerable attention because of their potential use
as biodegradable thermoplastics. By changing the carbon source and/or the
bacterial strain used in the fermentation process, it is possible to produce
biomaterials having properties ranging from stiff and brittle plastics to
rubbery polymers.
Poly(hydroxybutyrate) was first discovered in 1926 as a constituent of the
bacterium Bacillus megaterium. Since then, PHB and related PHAs have been
shown to occur in over 90 genera of bacteria. The majority of PHAs are
composed of R-(-)-3-hydroxyalkanoic acid monomers ranging from 3 to 14
6 Examples of Pathway Manipulations 245
R=hydrogen
R=methyl
R=ethyl
R=propyl
R--butyl
R=pentyl
R=hexyl
R=heptyl
R=octyl
R=nonyl
R O
I II
[-.O-CH-CH2-C-]x
3-hydroxypropionate (3HP)
3-hydroxybutyrate (3HB)
3-hydroxyvalerate (3 HV)
3-hydroxycaproate (3HC)
3-hydroxyheptanoate (3HH)
3-hydroxyoctanoate (3HO)
3-hydroxynonanoate (3 HN)
3-hydroxydecanoate (3HD)
3-hydroxyundecanoate (3HUD)
3-hydroxydodecanoate (3HDD)
n=3
n=4
FIGURE 6.13
O
II
[-O(-CH2-)nC-] x
4-hydroxybutyrate (4HB)
5-hydroxyvalerate (5HV)
Structures of major biological poly(hydroxyalkanoate)s.
carbons in length (Fig. 6.13). PHAs synthesized by bacteria can be broadly
subdivided in two groups: short-chain PHAs with C3-C5 monomers (e.g.,
Alcaligenes eutrophus) and medium-chain PHAs with C6-C14 monomers
(e.g., Pseudomonas oleovorans). Over 40 different PHAs have been character-
ized, with some polymers containing unsaturated bonds or other functional
groups (Steinbhchel, 1991).
PHB is the most widespread and thoroughly characterized PHA. Most
knowledge of PHB biosynthesis has been obtained from the bacterium
Alcaligenes eutrophus, which derives PHB from acetyl-CoA by the sequential
action of three enzymes (Fig. 6.14). The first enzyme of the pathway,
3-ketothiolase (or fl-ketothiolase), catalyzes the reversible condensation of
two acetyl-CoA molecules to form acetoacetyl-CoA. Acetoacetyl-CoA reduc-
tase then reduces acetoacetyl-CoA to R-(-)-3-hydroxybutyryl-CoA, which is
then polymerized by the action of PHA synthase to form PHB. Molecular
studies have revealed that the genes for these three enzymes are organized in
a single operon. PHA typically is produced as a polymer of 103-104
monomers, which accumulates up intracellularly as inclusion of 0.2-0.5/zm
in diameter. In A. eutrophus, PHB inclusions can typically accumulate to
80% of the dry weight when bacteria are grown in media containing excess
246 Metabolic Engineering
Propionic
Acid
ATP + CoASH
AMP + PPi
PropyionyI-CoA
3-ketovaleryl-OoA
., r e d u e t a s e :
~ . . . . . . . . . . . . . . . . . . . �9
Glucose
AcetyI-CoA (2x)
~ CoASH
acetoacetyl-CoA
/ ' - " NADPH + H §
NADP*
R - ( - ) - 3 - h y d r o x y v a l e r y I - C o A ~ R-(-)-3-hydroxybutyryI-CoA
CoASH "~--~*----"~ "" ...................... " ~"~.~ CoASH
FIGURE 6.14 Alcaligenes eutrophus pathways for PHB and P(3HB-3HV) synthesis.
carbon, such as glucose, but limited in one essential nutrient, such as
nitrogen or phosphate. Under these conditions, PHB synthesis acts as a
carbon reserve and an electron sink. When growth conditions are restored by
the addition of phosphate or nitrogen, PHB is catabolized to acetyl-CoA and
PHB returns to preinduction levels.
Induction studies on 3-ketothiolase and acetoacetyl-CoA reductase re-
vealed that both enzymatic activities increase markedly in response to
PHB-stimulating limitations. These experiments indicate that the PHB path-
way may exhibit a mode of transcriptional control that resembles these of
other metabolic pathways that are induced by environmental stress. Exam-
ples of such global regulatory networks include the heat shock regulon, the
pho regulon, and the carbon starvation regulon (see Section 5.4). Recently,
the A. eutrophus PHB biosynthetic genes (phaA, phaB, phaC) were cloned
and expressed in E. coli (Peoples and Sinskey, 1989; Schubert et al., 1988;
6 Examples of Pathway Manipulations 247
Slater et al., 1988) and various species of Pseudomonas (Timm and
Steinbi]chel, 1990). The A. eutrophus PHB pathway was found to be func-
tional in all recombinant strains, with PHB accumulation representing a
significant portion of the cellular dry matter when growth took place in
excess carbon source under nitrogen limitation. Interestingly, E. coli clones
produced PHB to approximately 50% of the level achieved in A. eutrophus
H16, while expressing reductase levels that were less than 2% of reductase
levels in A. eutrophus H16. Further subcloning identified two distinct forms
of A. eutrophus 3-ketothiolase, one believed to serve a biosynthetic role and
the other a catabolic role. The high levels of PHB achieved in certain
recombinant E. coli strains (up to 90% of cell dry weight) are indicative of
either a high degree of transcriptional versatility or a high degree of tran-
scriptional homology between the various strains. Another interesting result
was the fact that recombinant P. aeruginosa strains possess three different
pathways for the synthesis of poly(hydroxyalkanoate)s. When these cells are
grown on glucose, they accumulate a polymer consisting jS-hydroxybutyrate,
jS-hydroxydecanoate, and ]3-hydroxydodecanoate as the main constituents
and of ]3-hydroxyoctanoate and ]3-hydroxyhexanoate as minor constituents
(Timm and Steinbi]chel, 1990).Copolymers
At present, the PHA copolymer of greatest industrial interest is poly(3-
hydroxybutyric-co-3-hydroxyvaleric acid) [P(3HB-co-3HV)] due to its en-
hanced flexibility over the homopolymer [P(3HB)]. Addition of propionic
acid or valeric acid to the growth medium containing glucose leads to the
production of a random copolymer composed of 3-hydroxybutyrate and
3-hydroxyvalerate (Fig. 6.14). This random copolymer is currently produced
commercially under the brand name Biopol by Monsanto by fermentation of
the bacterium A. eutrophus on glucose and propionic acid. Incorporation of
various C3-C5 units in PHA is possible because of the broad specificity of the
bacterial enzymes involved in PHA synthesis. For instance, two 3-ketothio-
lases have been detected in A. eutrophus, which together accept from C4 to
at least C10 3-ketoacyl-CoAs, and the acetoacetyl-CoA reductase has been
shown to be active with C4-C6 3-ketoacyl-CoAs. By altering the intermediate
metabolism, Slater et al. have constructed an E. coli strain that produces this
copolymer at high titers (Slater et al., 1992). The strategy was based on
genetic elimination of the transcriptional regulation of E. coli genes of the
propionate pathway (constitutive expression), thus resulting in a strain that
can efficiently take up propionate and incorporate it into the copolymer
[P(3HB-co-3HV)]. Furthermore, this strategy introduced the ability to control
248 Metabolic Engineering
the ratios of the two polymers in P(3HB-co-3HV) by manipulating propionate
and/or glucose concentrations in the growth medium.
Substrates used to produce the biodegradable polymer poly(hydroxy-
butyrate) (PHB) in A. eutrophus include fructose, glucose, acetic or butyric
acid, and a mixture of H 2 and CO 2. A. eutrophus does not normally utilize
ethanol as a carbon source, but an ethanol-utilizing strain was engineered by
expressing the gene for ethanol dehydrogenase that converts ethanol to
acetaldehyde, which then enters the acetyl-CoA pool, a precursor of PHB.
More importantly, expression of the same gene allowed for the utilization of
propanol, which leads to the formation of the copolymer poly(hydroxy-
butyrate-valerate) (PHBV), a compound with a reduced melting point and an
improved polymer processibility compared with PHB (Alderete et al., 1993).
Up to 74% PHB by weight was obtained with 63 g L -1 dry cell mass. The
copolymer content increased with a higher fraction of propanol in the feed
and reached a maximum of 35.2 mol % from pure propanol.
A central issue of metabolic engineering in biopolymer production is the
maximization of polymer formation through the coordinated amplification of
the thiolase, reductase, and polymerase enzymes. Simple maximization of the
activity of all three is not the solution to this problem. PHB synthesis
depends on acetyl-CoA supply and NADPH availability. The first is maxi-
mized by increasing the rate of glycolysis; however, an increase in glycolytic
flux will reduce the flux into the pentose phosphate pathway and, hence,
NADPH generation. Therefore, maximization of PHA production is more an
issue of optimally balancing the flux distribution at the G6P branch point
rather than straightforward amplification of the three enzymes. This balance,
incidentally, may depend on the growth conditions, as the need for reduction
power and carbon may shift as cells pass from the growth to the production
phase.
Another important point to be made is that the relative activities of the
preceding three enzymes (thiolase, reductase, and polymerase) have a pro-
found impact on the quality of the product: Increasing the polymerase
activity while keeping the other two enzyme activities constant yielded PHB
with lower molecular weight, a counterintuitive result that can be explained
when one considers the actual mechanism of polymerization. Finally, the
production of the copolymer can be accomplished by feeding propionic acid
in the fermentation or by engineering the organism to provide its own supply
of propionic acid, such as through the threonine degradation pathway.
Plant Poly(hydroxyalkanoate)s
Recently the poly(hydroxyalkanoate) pathway has also been expressed in
crop plants, a very attractive system for such a purpose, with the potential
for producing large amounts of several chemicals at low cost. Synthesis of
6 Examples of Pathway Manipulations 249
PHB in plants initially was explored with the expression of PHB biosynthetic
genes of the bacterium A. eutrophus in the plant Arabidopsis thaliana (Poirier
et al., 1992). Although of no agricultural importance, A. thaliana was chosen
because of its extensive use as a model system for genetic and molecular
studies in plants, and because it is closely related to the oil-producing crop
rapeseed, a target crop for PHB production on an agricultural scale. Of the
three enzymes required for PHB synthesis, only 3-ketothiolase is present in
plants. In order to complete the pathway, A. eutrophus genes encoding the
acetyl-CoA reductase and PHA synthase were expressed in transgenic A.
thaliana, with activity found to be localized in the cytoplasm. This initial
attempt resulted in only 0.14% dry weight yield (approximately 2 orders of
magnitude below commercially attractive levels) and had an adverse effect on
cell growth.
A second generation of transgenic plants producing PHB proved to be
more successful. In plants, biosynthesis of fatty acids from acetyl-CoA occurs
in the plastid. The plastid is therefore a site of high carbon flux through
acetyl-CoA. This flux is particularly enhanced in the seeds of oil-accumulat-
ing plants, such as Arabidopsis, where up to 40% of the seed dry weight is
triglycerides. Furthermore, the plastid is also the site of starch accumulation,
and as such it can accommodate large amounts of inclusions without
disruption of organelle function. Expression of the PHB pathway in the
plastid recently has been demonstrated in transgenic A. thaliana (Nawrath
et al., 1994). Plastid expression was achieved by fusing the transit peptide of
the small subunit of ribulose bisphosphatase carboxylase to the N-terminus
of 3-ketothiolase and acetoacetyl-CoA reductase. The PHB content gradually
increased over the life span of the plant, reaching a maximum of 10 mg /g
fresh weight, representing approximately 14% dry weight. Thus, redirection
of the PHB from the cytoplasm to the plastid resulted in a 100-fold increase
in PHB production.
Fructan
Fructan is a poly(fructose) molecule naturally produced as a storage
compound in a limited number of plants and characterized by a low degree
of polymerization (5-60 units). Such polymers can be hydrolyzed enzymati-
cally or chemically to yield fructose, which is becoming an increasingly
popular sweetener in many food products. Because oligofructose molecules
are sweet, fructans themselves can be utilized directly as natural sweeteners.
Also, the human digestive system has no enzymes that can degrade the
[3(2 ~ 1) or fl(2 ~ 6) glycosidic linkages found in fructan, making this
sugar attractive as a low-calorie food ingredient. Besides plants, microorgan-
isms are capable of producing fructans of very high molecular weight
(> 100,000 units). For example, in Bacilli, Pseudomonas, and Streptococci,
250 Metabolic Engineering
extracellular fructosyltransferase converts sucrose to bacterial fructan, often
called levan. The main reaction for fructan biosynthesis is nGF (sucrose)
G-Fn (fructan) + n - 1G. For this purpose the SacB gene of B. subtilis, which
encodes the fructosyltransferase enzyme commonly known as levansucrase,
was modified and introduced into tobacco plants, resulting in transgenic
plants that can accumulate fructans. Production levels achieved a range from
3 to 8% dry weight, and the size andproperties of this fructan were found to
be similar to those of B. subtilis. An important feature of the recombinant
fructans is their stability in plants, which makes this work attractive for
applications in food and nonfood products.
Xanthan Gum
Xanthan is an extracellular polysaccharide produced by the Gram-negative
bacterium Xanthomonas campestris. Its unique rheological properties, such as
high viscosity and pseudoplasticity, account for its extensive use in a variety
of food and industrial applications. The chemical structure consists of a
cellulosic ]3(1 ~ 4)-glucose backbone with trisaccharide side chains com-
posed of two mannose residues and one glucuronic acid residue attached to
alternate glucose molecules on the backbone. Typically, the mannose sugars
are acetylated and pyruvylated at specific sites, but to various degrees. Many
of the genes involved in exopolysaccharide synthesis are often clustered. A
cluster of genes essential for xanthan synthesis has also been isolated in X.
campestris (Barrere et al., 1986). Recent studies have illustrated the potential
of recombinant DNA technology for altering the structure and properties of
xanthan gum. A plasmid containing several xanthan biosynthetic genes
increased the production of xanthan by 10%. Furthermore, by cloning and
overexpressing the gene for the enzyme ketal pyruvate transferase, the extent
of pyruvylation of the xanthan side chains was increased by up to 45%
(Harding et al., 1987). Conversely, using transposon mutagenesis, a strain
could be constructed that formed xanthan with a severely reduced pyruvate
content (Marzocca et al., 1991). Such studies are illustrative of the promise
of metabolic engineering in both the manipulation of product structure and
the elucidation of xanthan synthesis.
6 . 3 . 5 . BIOLOGICAL PIGMENTS
Indigo
One of the classic examples of metabolic engineering is the production of
indigo in genetically engineered E. coli carrying the naphthalene dioxyge-
6 Examples of Pathway Manipulations 251
nase gene from Pseudomonas putida, which catalyzes the final step in indigo
biosynthesis (Ensley, 1985) (see Fig. 6.15). Indigo, or indigotin, occurs as a
glucoside in many plants and has been used throughout history as a blue
dye. For the past century, indigo manufacturing has been carried out by
chemical synthesis leading to indoxyl, which is finally oxidized to indigo. By
using selective cultivation techniques, a soil organism (Pseudomonas indoloxi-
dans) was isolated in 1927 that could also decompose indole (Fig. 6.4) with
the formation of blue crystals later identified as indigotin. Even though
several other microbes were isolated that were also able to produce indigo
from indole, none was actually put to use in the large-scale microbial
synthesis of indigo due to (i) low availability of the precursor indole and
(ii) low activity of naphthalene dioxygenase (NDO), the final indigo biosyn-
thetic enzyme. Indigo production in these early strains required the cofeed-
ing of tryptophan or free indole, whose costs limited their use in commercial
processes.
Early attempts focused on enhancing the conversion of indole by overex-
pressing NDO from Pseudomonas putida. Superior enzyme activity was
obtained by combining the first four genes comprising the naphthalene
dioxygenase enzyme system into a multicopy plasmid under the control of
the strong ApL promoter. Further genetic manipulations were necessary to
improve the stability of heterologous NDO in E. coli. This work resulted in
highqevel expression of stable NDO that can be utilized for indigo biosyn-
thesis.
In the meantime, parallel efforts investigated synthesis of the indigo
precursor, indole, directly from glucose. Normally, indole is present in the
cell either in the form of indole 3-glycerol phosphate (IGP) or as a trypto-
phan moiety. In recombinant E. coli, tryptophan biosynthesis is carried out
by the five gene products of the tryptophan operon (Figs. 6.16 and 6.4). For
both bacteria and fungi, chorismate is the major branch point compound for
aromatic amino acid biosynthesis that includes phenylalanine, tyrosine, and
tryptophan. Also present in the cell is the enzyme tryptophanase, which
degrades tryptophan and releases indole. Early attempts, however, to stimu-
late indole production by overexpression of tryptophanase did not prove to
be very successful due to apparent limitation by the total tryptophan synthe-
sis flux. To correct for this, enzymes in the tryptophan synthesis pathway
were amplified. In particular, the trpB moiety was modified by site specific
mutagenesis which increased indole biosynthesis by more than an order of
magnitude (Murdock et al., 1993). An E. coli production strain was finally
developed that combined both enhanced indole production (mutated trpB)
and enhanced indole conversion to indigo (NDO), which can be used for
de novo indigo biosynthesis from glucose.
252 Metabolic Engineering
HC / CH i~ i fr Indole
Naphthalene
Dioxygenase
1
It HC~. .... ,+OH
HC "~c-----~
Indole dihydrodiol
J ?
HC /OH
HC / ~C-------~"
ii l rl HC~ ~C CH Indoxyl
i
Air
Oxidation
HC- 0
HC" ~C--------C ~ It i i Hc'Hc~C \ /c~
I Jl l o~c----c, c/H
Indigo
FIGURE 6.15 Indigo biosynthesis from indole by naphthalene dioxygenase (NDO). Indole
(see Fig. 6.4) is oxidized by NDO to form either an unstable dihydrodiol or indoxyl, which is
further condensed to form indigo by air oxidation.
6 Examples of Pathway Manipulations 253
6.3.6. HYDROGEN
Hydrogen could be the ultimate substitute for polluting and irreplaceable
fossil fuels for transportation, direct heating, or electricity generation. Be-
cause hydrogen combustion emits only water vapor and small quantities of
nitrogen oxides, hydrogen-fueled cars and other devices do not impact global
warming (carbon dioxide) and contribute only insignificantly to air pollution.
Prototype vehicles produced by manufacturers such as Mercedes Benz show
that hydrogen-powered cars can be practical in terms of performance,
comfort, and safety. Hydrogen is no more hazardous than methane or
gasoline, and it is, in fact, used routinely in industrial processes. Also, the
fact that the U.S. space program has relied on hydrogen as the fuel of choice
for nearly 40 years provides a knowledge base for extended uses of this fuel.
Conventional electrolysis techniques for extracting hydrogen from water,
though inexhaustible, still require slightly more energy than hydrogen would
yield upon combustion. Other means of hydrogen production have been
proposed, such as those utilizing wind or solar power (photovoltaic technol-
ogy), gasification, and pyrolysis.
Fermentation or enzymatic techniques are also currently being investi-
gated as potential biological routes to hydrogen production (Kitani and Hall,
1989; Taguchi et al., 1995) (Fig. 6.17). It is well-known that selected
microorganisms can efficiently produce hydrogen as an end product of
metabolism. Metabolic engineering techniques will prove very useful in
redirecting cellular metabolism toward hydrogen production above physio-
logical levels. Several genes involved in hydrogen synthesis have been
isolated thus far, such as the hydrogenase gene from Citrobacter freudii
cloned in E. coli (Kanamayia et al., 1988).
Recently, the possibility of in vitro hydrogen production has also been
investigated (Woodward et al., 1996). This system consists of two enzymes,
namely, glucose dehydrogenase (GDH) isolated from Thermoplasma aci-
dophilum and hydrogenase from Pyrococcus furiosus. GDH catalyzes the
oxidation of glucose to glucono-6-1actone, which further hydrolyzes to
gluconic acid using NADH or NADPH as a cofactor. Even though bacterial
hydrogenases rarely interact with NADPH becauseof its insufficiently low
potential, hydrogenases from P. furiosus and A. eutrophus have been shown
to use NADPH as an electron donor. Woodward et al. (1996) demonstrated
that the combination of GDH and hydrogenase was capable of hydrogen
production from glucose in vitro (Fig. 6.17). Stoichiometric yields of hydro-
gen were produced from glucose with continuous recycling of the cofactor
2 5 4 Metabolic Engineering
o % : H
!
H c / C ~ c H
II I CH2
HC..~c//C----O---C
I
I C- - -0
OH OH
trpEG ~
HC. /C( 0
HC//~ ~C ~ OH
i i
HC~. .C. HC/, ~H 2
trpD i V //o
HC.. _..C.~
HC ~/" ~C / ~'OH OH
I , ,,
HC<... /C. H --H% / HC-- HN~ 2~
"CHo/CH 2
trpF
HC. ~C: 0
HC//" ~C / "OH I It OH OH HC~-.. /C. l t
Hc ~ s ~ - - - ~ .=c- - -c H-c a - c . . - o - - ,r=o
I I HO trpC ~ HO OH
OH
s c ~ C ~ c . c c . - c . - c s r o - - ~ = o
it I II ' ' ' HC.. ~'C x OH HO OH HO
H c H C ~ c
11 I
HC~Hc~CX~HN~CH
- - C - - - - - - C H - - - C H - - C ~
II
Chorismate
Anthranilate
Phosphoribosyl
Anthranilate
CDRP
Indolglycerol
phosphate (IGP)
Tryptophan
FIGURE 16 Tryptophan biosynthesis. Structural gene designations: trpAB, tryptophan syn-
thase; trpC, indole glycerol phosphate synthase; trpD, anthranilate phosphoribosyl transferase;
trpEG, anthranilate synthase.
6 Examples of Pathway Manipulations 255
(at least 20 times). This newly discovered pathway seems to be an attractive
alternative for hydrogen production from renewable sources without the
immediate formation of waste gases such as CO 2 and CO. One limitation is
the need to identify further uses for the enormous amounts of gluconic acid
that would be produced as a byproduct of even a small-scale hydrogen
production plant. Future generations of such a process could involve immo-
bilized forms of these enzymes for continuous hydrogen synthesis.
Green algae, such as Chlamydomonas reinhardtii, could provide an appro-
priate mode of capturing light energy in fuel hydrogen (Cinco et al., 1993;
Lee et al., 1995). In a recent study (Greenbaum et al., 1995), it was shown
that mutation of the membrane-bound photosystem I reaction center does
not disable the complete photosynthesis. This original finding contradicts the
premise that both photosystems I and II are required for the conversion of
light energy to chemical energy, and it also doubles the maximum theoretical
conversion rate of light to chemical energy from 10 to 20%.
Renewable Resources
(Cellulose, Starch, Lactose)
Glucose
Glucose
Dehydrogenase
NADP §
HvdroQenase
NADPH
Glue~ lAcid ~'~
FIGURE 6.17 Conversion of renewable resources to hydrogen.
256 Metabolic Engineering
6.3.7. PENTOSES: XYLITOL
Xylitol, a sugar alcohol, is a good anticariogenic sweetener that does not
require insulin for its digestion by diabetics (Emodi, 1978). In nature xylitol
is found in certain fruits and vegetables in small amounts, making its
quantitative extraction difficult and uneconomical. Currently xylitol is manu-
factured chemically in alkaline conditions by catalytic reduction of xylose
derived from hemicellulose hydrolysates. Lately, much attention has focused
on the microbial production of xylitol from D-xylose. Xylitol has been
reported to be produced by yeasts, especially species of genus Candida, such
as C. pelliculosa (Nishio et al., 1989), C. boidinii (Vongsuvanlert and Tani,
1989), C. guillliermondi (Meyrial et al., 1991), and C. tropicalis (Gong et al.,
1981), Petromyces albertensis (Dahiya, 1991), by bacteria such as Enterobac-
ter liquefaciens (Yoshitake et al., 1973), Corynebacterium sp. (Yoshitake et al.,
1971), and Mycobacterium smegmatis (Izumori and Tuzaki, 1988). Yeasts
generally possess the first two enzymes needed for the metabolism of xylose:
xylose reductase (XR) and xylitol dehydrogenase (XDH) (Fig. 6.7). Efforts to
increase xylitol productivities and yields through culture optimization have
yielded moderate results (0.32-2.67 g L -1 h -I) (Horitsu et al., 1992). There-
fore, emphasis has been placed on genetic modifications that would enhance
both the yield and productivity of xylose conversion to xylitol.
In an initial study, the P. stipitis XR was chosen for the construction of
xylitol-producing recombinant yeasts (Hallborn et al., 1991). The choice was
based on the high specific activity of XR and the fact that this particular XR
uses both NADH and NADPH as cofactors (Verduyn et al., 1985). Due to the
lack of XDH (needed for NADH regeneration), the recombinant strain was
studied on medium containing both glucose and xylose. Xylose utilization
commenced after glucose exhaustion and proceeded to give about 97%
theoretical yield conversion of xylose to xylitol at a specific productivity of
0.08 g (gcells h) -1. A later study, aimed at expanding the substrate utilization
range of ethanologenic S. cerevisiae by overexpressing the P. stipitis genes for
XR and XDH, fortuitously resulted in the production of 35 g L -1 xylitol
(Tantirungkij et al., 1993).
6.4. IMPROVEMENT OF CELLULAR
PROPERTIES
This type of metabolic engineering is aimed at the organism as a whole; thus,
it is also referred to as cellular engineering. There are already a number of
successful applications of cellular engineering that involve a wide range of
6 Examples of Pathway Manipulations 257
organisms, from bacteria to animal cells. Such applications have been tar-
geted at improving specific growth rates and growth yields, providing resis-
tance to toxic compounds, improving the secretion of a specific product,
enhancing drought and salt tolerance of plant cells, and altering glycosyla-
tion sequences of recombinant polypeptides. It is an area of great challenges
and also vast opportunities.
6.4.1. ALTERATION OF NITROGEN METABOLISM
An early success of metabolic engineering was the alteration of the nitrogen
assimilation pathway of the methylotrophic bacterium Methylophilus meth-
ylotrophus to enhance the yield of single-cell protein (SCP). M. methylotro-
phus was the industrial choice for the production of SCP from methanol due
to its high carbon conversion efficiency, methanol tolerance, and nutritional
profile. However, the ammonia assimilation pathway of this organism has a
major drawback: it utilizes the glutamine synthase (GS) and glutamate
synthase system (GOGAT) that requires mol of ATP for every mole of
ammonia transported into the cell (see Section 2.4.1). In contrast, the
corresponding E. coli nitrogen assimilation pathway uses glutamate dehydro-
genase (GDH), which requires no ATP consumption (Fig. 6.18). M. meth-
ylotrophus probably uses the energetically suboptimal pathway for ammonia
assimilation because it evolved in an environment of low ammonia concen-
trations, as GS has a much higher affinity for ammonia than GDH. The
glutamate dehydrogenase gene (gdh) of E. coli cloned on a shuttle vector
was shown to complement gs mutants of M. methylotrophus (Windass et al.,
1980). As a result, the engineered organism exhibited higher methanol
conversion into cellular carbon, presumably because ammonia utilization is
more energy-efficient via GDH than via the coupled GS/GOGAT pathway.
The efficiency of carbon conversion was increased by 4-7%.
This work, which was one of the first industrial applications of metabolic
engineering, illustrated that the properties of organisms that evolved to
maximize the chances for survival in their natural habitat are not necessarily
optimal in the artificial environment of a large-scale bioreactor.
6 . 4 . 2 . ENHANCED OXYGEN UTILIZATION
A common engineering challenge in large-scale aerobic fermentations is
ensuring an adequate level of dissolved oxygen to achieve the desired cell
growth and productivity. Undermicroaerobic conditions that can arise, for
258 Metabolic Engineering
A. GS/GOGAT Pathway
NH4+ + ATP ADP + Pi
Synthase (GS) " ~
Glutamate Glutamine
Glutamate
,~nthase ( G ~
Glutamate 2-Oxoglutarate
+ NAD(P) + NAD(P)H
B. GDH Pathway
2-Oxoglutarate + NH4 § + NAD(P)H GDH .~ Glutamate + NAD(P)
FIGURE 6.18 Pathways of bacterial ammonia assimilation.
example, from nonideal mixing, elements of both respiration and fermenta-
tive metabolism are active and compete for accomplishing energy synthesis
and redox balance. Such oxygen fluctuations inevitably lead to undesirable
physiological events, such as the elicitation of global and specific oxygen
regulation responses that activate or repress key enzymes of central carbon
metabolism. Such survival responses usually are manifested as alterations in
growth rate and product formation (Onken and Leifke, 1989).
A recent finding that may in part alleviate the undesirable consequences of
oxygen fluctuation and hypoxic environments is the cloning of the
hemoglobin gene (vgb) from the bacterium Vitreoscilla (VHb) (Khosla and
Bailey, 1988a,b). Although its precise in vivo function has not yet been
established, VHb is thought to serve as an oxygen-binding protein enabling
Vitreoscilla to survive in microaerobic conditions characteristic of its natural
6 Examples of Pathway Manipulations 259
habitat. Recent studies indicate that VHb increases the number of protons
extruded per reduced oxygen atom across the cytoplasmic membrane and
enhances the ATPase-catalyzed ATP synthesis rate in microaerobic E. coli
(Chen and Bailey, 1994; Kalio et al., 1994) (see Section 2.3.3). Flux distribu-
tion analysis (Chapter 8) also revealed that VHb § cells have a smaller ATP
synthesis rate from substrate-level phosphorylation, but a larger overall ATP
production rate under microaerobic conditions (Tsai et al., 1995a-c). Fur-
thermore, results from cyo /cyd (aerobic/microaerobic terminal oxidases)
mutants suggest that the expression of VHb in E. coli increases the level and
activity of terminal oxidases and thereby improves the efficiency of microaer-
obic respiration and growth (Tsai et al., 1995). On-line culture NAD(P)H
fluorescence and redox potential measurements (CRP) suggested that Vhb
buffers intracellular redox potential perturbations caused by intracellular DO
fluctuations (Tsai et al., 1995).
Motivated by the hypothesis that this enzyme could possibly be beneficial
for growth in oxygen-limiting environments, the vgb gene was transferred in
various industrially important microbes. Heterologous expression of this
protein in a wide range of hosts has demonstrated that it elicits in vivo
effects of reduced oxygen starvation, improved cell growth, and enhanced
product formation (DeModena et al., 1993; Khosla and Bailey, 1988a,b;
Khosravi et al., 1990; Magnolo et al., 1991). For example, E. coli that carried
a single copy of this gene integrated on its chromosome synthesized total cell
protein more rapidly than an isogenic wild-type strain under oxygen-limiting
conditions. Furthermore, coexpression of VHb increases the expression of
cloned fl-galactosidase, chloramphenicol acetyltransferase (CAT), and a-
amylase by 1.5- to 3.3-fold relative to controls in oxygen-limiting E. coli
cultures. In other cases, expression of this protein achieved a 13-fold increase
in the production of an antibiotic in Streptomyces and a 1.2-fold increase in
the production of amino acids in Coryneform bacteria. Exogene, the company
founded to explore this technology, has also successfully expressed Vhb in
Penicillium and mammalian cells. Furthermore, in the field of bioremedia-
tion, transformation of Xanthomonas maltophilia with vgb resulted in an
enhanced efficiency of benzoic acid conversion to biomass (Liu et al., 1996).
6 . 4 . 3 . PREVENTION OF OVERFLOW METABOLISM
Currently, one of the current major technical challenges in recombinant
protein production processes employing E. coli is to maintain high intracel-
lular product levels at high cell concentrations. This dual goal is difficult to
achieve due to the accumulation of inhibitory culture byproducts. During
260 Metabolic Engineering
both aerobic and anaerobic growth, carbon and reductant fluxes are balanced
by the excretion of acidic byproducts, the most abundant of which is acetate.
This weak acid is a well-known growth inhibitor. Most importantly, it
reduces the cellular efficiency for the expression of recombinant products
and affects the quality of intracellular proteins, apparently by interfering with
disulfide bond formation.
Organic acids have been shown to influence cell growth at concentrations
that are low in comparison to inhibitory levels of mineral acids. The
undissociated forms of short-chain fatty acids produced intracellularly, such
as acetic acid, can freely permeate the cell membrane and accumulate in the
medium. Subsequently, a fraction of the undissociated acid that is present
extracellularly re-enters the cell, where it dissociates given the relatively
higher intracellular pH. This means that, in effect, weak acids act as proton
conductors (see Section 2.2.1 and Example 2.1). If this process continues
undiminished, the intracellular pH will approach the external pH and hence
the A pH component of the protonmotive force (see Section 2.33 and
Example 13.6) will collapse (Diaz-Ricci et al., 1990; Slonczewski et al.,
1981). In addition, a low extemal pH ( ( 5 ) can cause almost complete
growth stasis (without cell lysis) presumably due to the irreversible denatura-
tion of DNA and protein (Cherrington et al., 1991).
In addition to the preceding effects on cellular energetics, there are many
other factors contributing to the inhibitory nature of weak organic acids that
make minimization of acetate excretion a prerequisite for optimizing the
production yields of recombinant processes (Yee and Blanch, 1992; Zabriskie
and Arcuri, 1986). The chemostat data of Jensen and Carlsen (1990), who
studied the effects of acetate on the production of human growth hormone in
E. coli, clearly illustrate the significance of acetate in this recombinant
system. By varying the amount of acetate in the feed, it was determined that
acetate levels of 40 mM reduce recombinant protein yields by approximately
35% without having any effect on the biomass yield. This result agrees with
the general observation that the acetate threshold that influences recombi-
nant protein yields is usually lower than that that causes notable growth
inhibition. In the same study, increasing the acetate level to 100 mM caused
a reduction in biomass yield by more than 70%, whereas recombinant
product yield declined by a factor of 2. Several other investigators have
implicated acetate as an important factor in the deterioration of recombinant
process productivities (Brandes et al., 1993; Brown et al., 1985; Curless
et al., 1988; Luli and Strohl, 1990; Starrenburg and Hugenholtz, 1991).
It is widely accepted that acetate excretion results from an imbalance
between the glycolytic flux and the cell's actual requirements for metabolic
precursors and energy. Pyruvate, which is the end product of glycolysis as
6 Examples of Pathway Manipulations 261
well as the precursor of acetate, provides a suitable junct ion for effecting
acetate accumulation. The strategy involves the introduction of a heterolo-
gous enzyme to catalyze the redirection of surplus carbon flux to a less
harmful byproduct than acetate. The B. subtilis acetolactate synthase (ALS)
enzyme was selected for this purpose on the basis of the fermentation
characteristics of this group of microorganisms (Johansen et al., 1975). Table
6.6 compares the amounts of fermentation byproducts excreted by mixed
acid producers, such as E. coli,with those of members of the butanediol
family, namely, Bacillus subtilis and Aerobacillus polymyxa. Evidently, mem-
bers of the second group form very small amounts of acids compared with
E. coli while they convert glucose primarily to the neutral compound
2,3-butanediol.
As shown in Fig. 6.19, butanediol producers normally have two distinct
enzymes that convert pyruvate to acetolactate: the "pH 6" acetolactate
synthase (ALS) and the acetohydroxy acid synthetase (AHAS). AHAS, an
anabolic enzyme found in many microorganisms, also catalyzes the initial
steps from pyruvate in the formation of the branched-chain amino acids
valine, leucine, and isoleucine. AHAS is also a flavoprotein that is regulated
by end product feedback inhibition, such as by valine. On the other hand,
ALS does not require FAD for activity, nor is it inhibited by the presence of
branched amino acids (St/Srmer, 1968).
The alsS gene from B. subtilis encoding the acetolactate synthase enzyme
was successfully expressed in E. coli (Aristidou et al., 1994a,b). This enzyme
TABLE 6.6 Comparison of Mixed Acid and Butanediol Fermentations a
Mixed Acid Butanediol
Products b E. coli P. formicans B. subtilis A. polymyxa
2,3-Butanediol 0.26 - 54.60 65.1
Acetoin 0.19 - 1.56 2.8
Glycerol 0.32 - 56.80 -
Ethanol 50.5 64.0 7.65 66.2
Formate 86.0 105.0 1.32 -
Acetate 38.7 62.0 0.16 2.9
Lactate 70.0 43.0 17.61 -
Succinate 14.8 22.0 1.08 -
CO 2 1.75 - 117.8 199.6
H 2 0.26 - 0.16 70.9
C recovery (%) 94.7 96.0 98.0 101.6
O/R balance 0.91 1.02 0.99 0.99
a From Wood, 1961.
b mmoles/100 mmoles of fermented glucose.
262 Metabolic Engineering
Pyruvate
AHAS
+ a-ketobutyrate ~ a-aceto a-hydroxybutymte ~ ~ ~ isoleucine
ALS
+ Pymvate ~ ot-acetolactatc ~ ~ ~ valinc or. /
diacctyl ~ALDC
N A D H ~ ~
NAD + acetoin
AIR ~ N A D H
2,3-butanediol leucine
FIGURE 6.19 Comparison of ALS and AHAS enzymes and their roles in branched-chain
amino acid synthesis and butanediol formation. Abbreviations: ALS, acetolactate synthase;
AHAS, acetohydroxy acid synthetase; c~-ALDC, a-acetolactate decarboxylase; AR, acetoin reduc-
tase (or 2,3-butanediol dehydrogenase); DAR, diacetyl reductase.
acts at the pyruvate branch point, redirecting excess carbon flux away from
acetate and toward the noninhibitory byproduct a-acetolactate. Characteriza-
tion of the resulting strain indicated that acetate excretion can be maintained
below 20mM even in dense cultures employing rich glucose medium. More-
over, the engineered strain is a more efficient host for the production of
recombinant proteins (Aristidou et al., 1994): the volumetric expression of
recombinant fl-galactosidase was found to increase by about 50% in batch
cultivations and by about 220% in high cell density fed-batch cultivations.
These results demonstrate the successful application of metabolic engineer-
ing for the improvement of cellular characteristics.
6 . 4 . 4 . ALTERATION OF SUBSTRATE UPTAKE
Nutrient uptake by transport through the cell membrane is an important task
of all living organisms. In addition to free diffusion, transport mechanisms
fall into three general categories, namely, (i) facilitated diffusion, (ii) group
translocation, and (iii) active transport (see Section 2.2). Hexoses, like
glucose, mannitol, and fructose are primarily transported into the cell by the
phosphoenolpyruvatedependent carbohydrate:phosphotransferase system
(PTS), which is a group translocation process (Dills et al., 1980; Postma
6 Examples of Pathway Manipulations 263
et al., 1993; Saier et al., 1988). The overall process catalyzed by PTS can be
summarized as
- (PEP)I N - (carbohydrate)ouT + (pyruvate)i N + (carbohydrate-Pi)iN--0
regardless of the carbohydrate or microorganism. The PTS accomplishes both
the translocation and phosphorylation of the carbohydrate in a series of steps
that involves a number of cytoplasmic as well as membrane-bound proteins
(Section 2.2.3).
In addition to serving an important role in sugar uptake, PEP is also an
essential precursor for several specialty chemicals, including aromatic amino
acids, indigo, enterobactin, and melanin. Thus, by providing a non-PTS sugar
uptake alternative, one should, in principle, save 1 mol of PEP for every
glucose consumed. Some of the approaches utilized to improve the availabil-
ity of PEP for biosynthetic purposes include the use of non-PTS carbon
sources, pyruvate recycling to PEP by PEP synthase overproduction (Patnaik
and Liao, 1994; Patnaik et al., 1995), and inactivation of pyruvate kinase
(Mori et al., 1987). A major breakthrough in this area resulted from deletion
of the glucose PTS genes ptsH, ptsI, and crr of an E. coli strain so that
glucose could no longer be transported into the cell by the PEP-consuming
system (Flores et al., 1996). "Revertant" strains subsequently were selected in
chemostat cultures, which were later characterized to possess a galactoseper-
mease gene that is able to transport glucose efficiently. The stable, rapidly
growing engineered strain NF9 subsequently was used in a genetic back-
ground of elevated DAHP synthase (the enzyme that condenses PEP and
erythrose-4-P into DAHP), where it was illustrated that PEP saved during
glucose transport was redirected into the aromatic pathway.
6 . 4 . 5 . MAINTENANCE OF GENETIC STABILITY
During recent years, the use of plasmid cloning vectors as carriers of genes
whose products are of scientific or commercial interest has become common.
Some of the desirable characteristics of a plasmid vector include small size,
high or controllable copy number depending on the application, strong and
controllable promoter for high-level gene expression, and, most importantly,
structural and genetic stability. Genetic instability is a primary impediment
to industrial utilization of recombinant microorganisms. There can be vari-
ous reasons leading to such instability, and in general expression vectors that
are most effective in directing protein production are usually more unstable.
This is perhaps a consequence of the elevated metabolic burden imposed on
the cell and can lead to a plasmid-free population within a few generations.
264 Metabolic Engineering
In addition to segregational instability, structural instability through homolo-
gous recombination events can lead to plasmid derivatives that no longer
produce the desired product.
Prevention of segregational and structural instability has been the subject
of intense research in the last 15-20 years. Structural instability generally is
minimized by deletion of the recA gene from the host cell. This deletion
should severely limit homologous recombination between extrachromosomal
and chromosomal DNA. In their early work, Csonka and Clark (1979)
showed that the A(srl_recA)306 mutation reduced the rate of recombination
by a factor of 36,000. Additionally, Laban and Cohen (1981) have shown that
a recA point mutation lowers the frequency of recombination events within a
plasmid by 100-fold. Segregational instability, on the other hand, is a more
challenging issue that requires more elaborate solutions.
Several strategies to obtain genetically stable plasmids have been devised.
The most primitive solution seems to be the addition of antibiotics to the
growth medium thus selecting for cells harboring the plasmid vector with
antibiotic resistance. This approach has a number of obvious drawbacks, e.g.
the use of costly and contaminating antibiotics. A more pragmatic approach
was suggested by Skogman and Diderichsen (Diderichsen, 1986; Skogman
et al., 1983). These authors used a special host-vector system where a specific
chromosomal mutation giving rise to an auxotrophic host is complemented
by a corresponding functional gene on the plasmid vector of interest.Although a powerful approach, it has applicability limitations due to the
requirement of special host strains carrying specific mutations.
A technique of wider applicability involves the hok/soh locus (formerly
parB) of plasmid R1. This system was discovered through its ability to
mediate efficient stabilization of a variety of plasmids in Gram-negative
bacteria (Gerdes, 1988; Gerdes et al., 1986). Later it was illustrated that the
increased plasmid maintenance was a consequence of the selective killing of
cells that at the point of division lose the hok/sok-bearing plasmid. The
hok/sok locus codes for two RNAs, Hok (host killing) mRNA and Sok
(suppression of killing) RNA. The hok product is a potent cell-killing agent
that destroys bacterial cells from within by damaging the cell membrane. The
sok product is a trans-acting antisense RNA (Sok-RNA, see Box 6.3) that
represses hok gene expression at a post-transcriptional level. The rapid and
selective killing of plasmid-free segregates can be explained on the basis that
hok mRNA is extremely s table ( t l / 2 ~, 20 min), whereas sok mRNA decays
very rapidly. Hence, in a plasmid-carrying cell, the Sok RNA prevents
synthesis of the Hok protein and the cell remains viable. On the other hand,
when a plasmid-free segregate appears, the unstable Sok RNA molecules
decay, thereby rendering the stable Hok RNA accessible to translation.
6 Examples of Pathway Manipulations 265
BOX 6.3
Antisense RNA
Natural antisense RNAs are small (15-50 nucleotides), untranslated,
regulatory RNA molecules that inhibit the function of their target
RNAs, to which they are complementary. Antisense RNAs regulate such
diverse functions as plasmid replication, conjugation, and maintenance,
transposition, and lysis-lysogeny decisions in bacteriophages. Antisense
RNAs recognize their target RNA via an initial reversible contact
between a single-stranded loop in the antisense RNA and a complemen-
tary loop in the target RNA. The initial contact usually is followed by
formation of a thermodynamically very stable duplex between the two
RNA molecules.
Antisense RNAs may inhibit the function of their target RNAs by
several different mechanisms. For example, the transposable gene of
Tnl0 is regulated by RNA-OUT, an antisense complementary to the
translational initiation region (TRI) of the transposable mRNA. In this
case, it has been shown that hybridization of the antisense RNA to the
mRNA physically blocks entry of the ribosome at the transposable gene
and, thus, is an example of direct target RNA inhibition mediated by an
antisense RNA.
Indirect target RNA inhibition by antisense RNAs has been described
in several cases. The classic example is perhaps the replication control
circuit of plasmid ColE1. The antisense RNA (RNA I) interacts with
RNA II, the primer of replication initiation, and thereby induces a
secondary structure change several hundred nucleotides downstream
from the hybridization site. The change in secondary structure prevents
the ribonuclease (RNase) H-dependent cleavage of the primer RNA that
is a prerequisite for replication initiation.
266 Metabolic Engineering
Consequently, the Hok protein is synthesized, thus leading to the death of
the plasmid-free segregates.
6.5. XENOBIOTIC DEGRADATION
Natural processes, both biological and geochemical, produce enormous
amounts of diverse organic compounds, and, over the eons of evolution,
different microbes have developed the ability to degrade nearly all such
natural compounds as a source of carbon and/or energy. However, many of
the tens of thousands of artificial organic compounds produced by humans
for industrial or agricultural purposes have no apparent counterparts in the
microbial world. Such synthetic novel compounds are called xenobiotics,
from the Greek word xenos, which means "foreign." Xenobiotics (such as the
PCBs discussed later) are stable compounds that are also fat-soluble; thus,
they become increasingly concentrated as they travel up the food chain.
Xenobiotic degradation is a rapidly developing area that holds great
opportunities for metabolic engineering. It involves primarily the utilization
of genetically engineered microbes for the degradation of pollutants and is
commonly known as bioremediation. Bioremediation currently is being used
to decrease the organic chemical waste content of soils, ground water, and
effluent from chemical plants and food processing and oil sludge from
petroleum refineries.
Early attempts for the isolation of xenobiotic-degrading organisms focused
on chemostat selection methods. This technique is based on inoculating
chemostats with microbial samples from various toxic waste dump sites and
selecting for desirable strains. By using this approach, it was possible to
isolate a Pseudomonas strain capable of degrading halogenated compounds,
such as the herbicide 2,4,5-trichlorophenoxyacetic acid (Kilbane et al.,
1983). The isolated strain was successful in removing 98% of the pollutant
from soil within a week, after which it died rapidly, as it was unable to
compete with endogenous strains.
In addition to chemostat selection, rational approaches for designing
useful catabolic pathways have also been reported in the last decade, some of
which are discussed next.
6.5.1. POLYCHLORINATED BIPHENYLS (PCBs)
PCBs are a class of 209 distinct man-made compounds carrying 1-10
chlorines attached to a biphenyl (Fig. 6.20). From 1929 to 1978, approxi-
mately 700,000 tons of PCBs were produced, with several hundred thousand
6 Examples of Pathway Manipulations 267
3 2 4(--)/
5 6
2~ 3'
6' 5'
4 ~
Biphenyl
FIGURE 6.20 General polychlorobiphenyl (PCB) structure. Numbers indicate possible chlori-
nated sites. Empirical formula: C12H12_nCI n where n = 1-10.
tons of that released into the environment. Although there are naturally
occurring microbes that can oxidize the mono-, di-, and trichlorinated PCBs,
this is not the case for the higher chlorinated PCBs. Cometabolism is
probably the most common method of PCB degradation in nature, but this is
a slow process as the initial transformations do not provide carbon or energy
for growth.
The genes for PCB-degrading enzymes ( bph A, -B, -C, and-D) have been
isolated from the Pseudomonas strain LB400 (Fig. 6.21) and subsequently
overexpressed in an E. coli strain. The PCB-degrading ability of the recombi-
nant strain was comparable to that of LB400 in terms of both specificity and
the extent of degradation.
By introducing specific genes into Pseudomonas strains it has been possi-
ble to construct strains that are able to degrade mixtures of aromatic
compounds, such as 3-chloro- or 4-methylbenzoate, which are frequently
present in industrial wastes (Rojo et al., 1987). Another interesting enzyme
system for the degradation of various aromatic compounds is the ligninase of
the white rot fungus Phanerochaete chrysosporium, which can decompose a
broad range of xenobiotics, even including such diverse structures as ben-
zopyrene and triphenylmethane dyes (Bumpus and Brock, 1988).
6 . 5 . 2 . BENZENE, TOLUENE, AND p - XYLENE
MIXTURES (BTX)
This group of petroleum-derived contaminants are water-soluble and, thus,
can pollute water resources, posing serious health threats to humans. A
number of studies have focused on the development of genetically engi-
neered strains that would convert such pollutants, either individually or
268 Metabolic Engineering
HC
.~Jr162 c ~ ~c~ rl rr HCJ /CH HC~ CH
cI~HC ~" HC - ~ ~ bphA
HO
H~ T OH ~cYC~__c/C'~c~H
I1 I It I
cI~HC /2"CH HO.. ~CH HC l bphB u HO
I OH
Hc~C-~c c r ~
II I I II HC~r ~CH HC~.. ./C H
C HC ~
C1 I bphC
HO
/CH % \ //0/OH C\
HC ~ "~C---/C C
II I I II HC..~/~CH HC~. //CH
S CH HC" CI ~ bphD
0
tl
HC IC..
HC" ~'C / ~OH
II I HC ~cu
C17 Hc
FIGURE 6.2.1 Degradation of chlorobiphenyl by enzymes of the 2,3-dioxygenase pathway of
Pseudomonas strain LB400. Gene designations: bphA, bipheny| 2,3-dioxygenase; bphB, dihydro-
dio| dehydrogenase; bphC, 2,3-dihydroxybipheny] dioxygenase; bphD, 2-hydroxy-6-oxo-6-phen-
ylhexa-2,4-dienoic acid hydrase.
6 Examples of Pathway Manipulations 269
collectively, to less harmful compounds, such as pyruvate. Two such exam-
ples are discussed next.
TOL (pWW0) Catabolic Plasmid
Catabolic plasmids typically contain a complete set of genes required for a
certain degradative pathway and are particularly ubiquitous in Pseudomonas.
They are self-transmissible, and many have a broad host range making them
particularly useful in nature, where such catabolic pathways can be made
available to different species. Many catabolic plasmids isolated thus far carry
pathways for the degradation of aromatic compounds, presumably because
the benzene ring is second in natural abundance to glucose as a building
block.
The TOL (pWW0) catabolic plasmid from Pseudomonas putida has been
shown to confer on its host the capacity to degrade toluene, as well as m-
and p-xylene and other benzene derivatives. The genes are organized in two
operons (see Table 6.7 and Fig. 6.22): (1) xylCAB, which encodes the
degradation of toluene and xylenes to benzoate and toluates, respectively
(Upper pathway), and (2) xylXYZLEGFJKIH, which encodes the degradation
of benzoate and toluates to acetaldehyde and pyruvate (Lower pathway). The
branching at 2-hydroxymuconic semialdehyde broadens the substrate range
TABLE 6.7 Genes and Corresponding Products of the TOL Catabolic Plasmid pWYq0
Gene Enzyme or function
Upper pathway operon
xylA
xylB
xylC
Lower pathway operon
xylX, Y,Z
xylE
xylF
xylG
xylH
xylI
xylJ
xylK
xylL
Regulation
xylR
xylS
Xylene oxygenase
Benzyl alcohol dehydrogenase
Benzaldehyde dehydrogenase
Toluene dioxygenase
Catechol 2,3-dioxygenase
2-Hydroxymuconic semialdehyde hydrolase
2-Hydroxymuconic semialdehyde dehydrogenase
4-Oxalocrotonate tautomerase
4-Oxalocrotonate decarboxylase
2-Oxopent-4-enoate hydratase
2-Oxo-4-hydroxypentenoate
Dihydroxycyclohexadiene carboxylase dehydrogenase
Transcription regulation
Transcription regulation
2 7 0 Metabolic Engineering
CH3 CH2OH CliO COOH
HOOC.,..~ j O H
C ~
Toluene Benzyl alcohol Benzaldehyde Benzoic acid
O
~c~~
.o~..
OH 002
,OH
Catechol x~~
OH OH
COOH xyJG ~ ~COOH
L~,,./COOH
2-Hydroxymuconic 2-Hydmxy-2,4-
Semialdehyde hexadlene-1,6-
d l o x a n e
xylF ~ HCOOH
o
CO2
COON
r
xj41
1,2-Dihydroxycyclo-3,5-
hexadlene carboxylate
, I
xylH ~ C O O H
--------~ I ~ I ~ C 0 0 H
2-Oxo-3-hexene-
1,6-dioxane
2-Oxo-4-pentenoate
O ~
"~COOH +
Pyruvtc Acid
, , O H
Acetaldehyde
FIGURE 6.22 Toluene degradation by Pseudomonas putida recombinant strain mt-2.
6 Examples of Pathway Manipulations 271
that can be degraded by such pathway. For example, toluate is degraded by
the xylF branch, whereas benzoate and p-toluate are degraded by the
xylGHI branch.
Genetic engineering techniques have been applied in order to extend the
range of substrates that an organism can utilize. For example, P. putida
carrying the TOL plasmid is able to grow on a variety of alkylbenzoates, such
as benzoate, 3- and 4-methylbenzoate (3MB and 4MB), 3,4-dimethylbenzoate
(34DMB), and 3-ethylbenzoate (3EB), but not on the closely related com-
pound 4-ethylbenzoate (4EB). A series of experiments to explain this con-
cluded that (1) 4EB did not activate a key regulatory protein (xylS), and as a
result, genes of the benzoate catabolic pathway (xylABC, Fig. 6.22) remained
uninduced, and (2) the native enzyme catechol 2,3-dioxygenase (xylE, Fig.
6.22) was unable to use 4-ethylcatechol as substrate (but could use 4-methyl-
catechol, for instance) (Ramos and Timmis, 1987). Thus, 4EB degradation by
this pathway would require (1) a xylS regulator with a broader effector
specificity and (2) generation of catechol 2,3-dioxygenase mutant enzymes
capable of degrading 4-ethylcatechol. Both strategies have been pursued,
leading to the isolation of mutated xylS and xylE genes, whose products
behave in accordance with the preceding requirements (xylS' and xylE').
P. putida cells containing a TOL plasmid with both the xylS' and xylE' genes
were indeed able to grow on all the usual substrates mentioned previously as
well as on 4EB.
Simultaneous Biodegradation of BTX Mixtures
Because BTX compounds typically are discharged into the environment as
a mixture of the three aromatics, effort was placed on developing a strain
that could simultaneously degrade all three pollutants (Lee et al., 1994). To
this end, a P. putida strain was developed that contains two catabolic
pathways: the TOL pathway discussed previously and the TOD pathway.
Enzymes of the TOD pathway act on all three aromatics; however, the end
products of this pathway (benzene-, toluene-, and p-xylene-cis-glycol) can-
not be metabolized further. On the other hand, enzymes of the TOL pathway
can act only on toluene and xylene; however, its end products can be utilized
for energy and carbon intermediates. The general form of the combined
catabolic pathways is summarized in Fig. 6.23.
The original strain was constructed by including all enzymes of TOL and
TOD. Characterization studies indicated, however, that this strain accumu-
lated significant amounts of TOD end products, i.e., benzene-, toluene-, and
p-xylene-cis-glycols. In order to alleviate this, the final step of the TOD
pathway, toluene-cis-glycol dehydrogenase, was blocked as indicated in
Fig. 6.23. The final strain was able to degrade all three aromatics at
2 7 2 Metabolic Engineering
Benzene Toluene Xylene
CH3
CH3 !
.
o
' * * o �9 ~ . - . . =
\ .....-- . ................. j " o.~
, ~149176 . . . . o . . . . ~176176 "'~ �9 ~" Xylene oxygenase
�9 1 ~ ..... Toluene dioxygenase .... Benzyl alcohol dehydrogenase ... Benzaldehyde dehydrogenase
�9 �9 _ . .
. - " '... OH ~ ~OOH
OH ~li COOH
~ p-xylene-cis-glycol
o~,ec~o, . , ,., OH~
uene-cis-gJyco, ..'" J Toluate dioxygenase I
HOOC ~ OH
HOO OH
OH
benzoate- c/s-glycol CH3
p-toluate-cis-glycol
J
Toluene-cis-glycol dehydrogenase I ~ Toluate-c/s-glycol dehydrogenase
I I Unidentified E~ 'OH { ~ ~ ' ~ ~
Intermediates
IT Ring cleavage, further I degradation leading to ~r CA Cycle IntermediatesJ
. •
.... $ . /" ~
{ Catechol 2,3-dioxygase, xylE I
CH3
FIGURE 6.23 Metabolic pathways for the degradation of benzene, toluene, and p-xylene in
Pseudomonas putida. Dashed lines represent the TOD pathway and solid lines the TOL pathway
(see also Table 6.7 and Fig. 6.22). (-) represents genetic blockage in the TOD pathway
(toluene-cis-glycol dehydrogenase).
6 Examples of Pathway Manipulations 273
significant rates: 0.27, 0.86, and 2.89 mg (mg biomass h) -1 for benzene,
toluene, and p-xylene, respectively.
This is an interesting illustration of how existing catabolic plasmids (that
constitute complete pathways) can be combined judiciously to generate new
organisms with novel properties, such as wider substrate utilization range or
better product formation.
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